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Journal of Virology, November 2002, p. 11033-11041, Vol. 76, No. 21
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.21.11033-11041.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Productive Infection of Plasmacytoid Dendritic Cells with Human Immunodeficiency Virus Type 1 Is Triggered by CD40 Ligation

Lawrence Fong,1* Manuela Mengozzi,2 Nancy W. Abbey,3 Brian G. Herndier,3 and Edgar G. Engleman1

Department of Pathology,1 Department of Genetics, Stanford University School of Medicine, Stanford, California 94304,2 Department of Pathology, University of California, San Francisco, California 941103

Received 18 March 2002/ Accepted 19 July 2002


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ABSTRACT
 
Immature plasmacytoid dendritic cells are the principal alpha interferon-producing cells (IPC), responsible for primary antiviral immunity. IPC express surface molecules CD4, CCR5, and CXCR4, which are known coreceptors required for human immunodeficiency virus (HIV) infection. Here we show that IPC are susceptible to and replicate HIV type 1 (HIV-1). Importantly, viral replication is triggered upon activation of IPC with CD40 ligand, a signal physiologically delivered by CD4 T cells. Immunohistochemical staining of tonsil from HIV-infected individuals reveals HIV p24+ IPC, consistent with in vivo infection of these cells. IPC exposed in vitro to HIV produce alpha interferon, which partially inhibits viral replication. Nevertheless, IPC efficiently transmit HIV-1 to CD4 T-cells, and such transmission is also augmented by CD40 ligand activation. IPC produce RANTES/CCL5 and MIP-1{alpha}/CCL3 when exposed to HIV in vitro. IPC also induce naïve CD4 T cells to proliferate and would therefore preferentially infect these cells. These results indicate that IPC may play an important role in the dissemination of HIV.


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INTRODUCTION
 
Plasmacytoid dendritic cells (PDC) are unusual in that they participate in both innate and adaptive immunity (8, 33). In an immature state, these bone marrow-derived cells are the primary cells that produce alpha interferon (IFN-{alpha}) in response to viral infection. Mature IPC, also called PDC or DC2, function as antigen-presenting cells capable of inducing immunity in naïve allogeneic T cells similar to classical myeloid DC (MDC). Viral infection in particular triggers IPC to induce Th1 immunity (7, 18). In contrast to MDC, which are derived from a committed MDC precursor or granulocyte-monocyte precursor, IPC express high levels of the interleukin 3 (IL-3) receptor (CD123) and lack myeloid markers such as CD11c (8, 15, 25, 31). A recent study suggests that IPC may be of lymphoid origin (35). Originally termed "plasmacytoid T cells" because of their unique plasma-cell-like morphology and their localization in the T-cell zones of lymph nodes (LN) (38), these cells have been identified in blood, tonsil, and LN (15, 17, 25). IPC precursors require IL-3 for survival and an activation stimulus, such as CD40 ligand (CD40L), for maturation into PDC (15).

While the susceptibility of DC to human immunodeficiency virus (HIV) has been extensively studied, most reports have focused on either MDC or bulk DC populations that would represent a mixture of MDC and IPC (4, 20, 26, 28). The susceptibility of IPC to HIV infection has been difficult to examine because of their scarcity, representing only 0.1% of peripheral blood mononuclear cells (PBMC). Nevertheless, IPC have been identified in sites, such as blood and tonsil (15), where HIV is transmitted and, therefore, could also be exposed to virus during infection. Moreover, a progressive loss of IPC in the blood has been observed in HIV-infected patients, suggesting an interaction between IPC and HIV (9, 34). A preliminary report suggests that IPC may be susceptible to HIV, although neither the molecular basis for infection nor the effect of IPC maturational status on infectivity was explored (27).


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MATERIALS AND METHODS
 
Flow cytometry. Cells were stained with the following antibodies: Per-CP labeled antibodies against HLA-DR; allophycocyanin (APC)-labeled anti-CD3, -4, -14, -19, and -56; FITC-labeled anti-CD11c, -CD62L, -CCR5, and -CXCR4 and annexin V; and phycoerythrin-labeled anti-CD123 (Becton Dickinson, San Jose, Calif.). Fluorescein isothiocyanate (FITC)-labeled anti-DC-SIGN was obtained from eBioscience (San Diego, Calif.). FITC-labeled anti-p24 antibody was obtained from Beckman-Coulter (Brea, Calif.). Propidium iodide (PI) was purchased from Sigma (St. Louis, Mo.). Prior to antibody staining, cells were preincubated for 15 min in Dulbecco's phosphate-buffered saline containing 0.01% sodium azide, 2% fetal calf serum, and Fc-blocking human immunoglobulin G at a concentration of 1 mg/ml (Sigma). Cells were then stained with the corresponding antibodies for 30 min at 4°C. Cells were then washed and analyzed with a four-color FACSCalibur (Becton Dickinson). For intracellular staining, cells were stained with surface marker antibodies, fixed with 4% paraformaldehyde, permeabilized with 0.5% saponin (Sigma), and stained with anti-p24 antibody. For annexin V staining, cells were stained with surface antibody, washed, and then stained with annexin V-FITC and PI in 2.5 mM CaCl2 and 140 mM NaCl buffer for 30 min and then analyzed.

In vivo expansion of human DC. HIV-seronegative study patients participating in a cancer vaccine trial, who provided signed informed consent that fulfilled Institutional Review Board guidelines, received 10 daily subcutaneous injections of Flt3L (Immunex, Seattle, Wash.) at a dose of 20 µg/kg of body weight/day to a maximum daily dose of 1.5 mg/day. Patients then underwent leukapheresis, and PBMC were obtained with Ficoll separation of the apheresis product.

Chemotaxis assays. Chemotaxis assays were performed as previously described (24). Briefly, 106 PBMC in 100 µl of complete medium (CM) comprised of RPMI 1640 (BioWhittaker, Walkersville, Md.) with 10% human AB+ serum supplemented with IL-3 at a saturating concentration of 10 ng/ml (Peprotech, Rocky Hill, N.J.) were added to each 24-well 5-µm-pore-size transwell insert (Corning, Acton, Mass.) with 600 µl in the lower chamber, with or without chemokine. SDF-1/CXCL12 and MIP-1{alpha}/CCL3 (R&D Systems, Minneapolis, Minn.) were added at concentrations ranging from 1 to 1,000 ng/ml. After 3 h of incubation at 37°C, the transwell inserts were removed. The cells in the lower chamber were collected, stained with antibody, and enumerated by collecting events for a fixed time (60 s) on a FACSCalibur. By counting a 1:5 dilution of input cells, the absolute number of cells that transmigrated could be obtained. To determine which subsets of DC were migrating, cells were stained and analyzed by four-color flow cytometry as described. Results are expressed as the percentage of input DC per transwell. Standard deviations are shown for three replicate transwells. Data are representative of three separate experiments.

Human DC isolation. PBMC were labeled with mouse monoclonal antibodies to human CD3, CD14, CD19, and CD56 and washed. Antibody-labeled PBMC were then depleted with anti-mouse immunomagnetic beads (Dynal, Lake Success, N.Y.). This negative selection was followed by positive selection with immunomagnetic beads for major histocompatibility complex class II (MHC-II) (Miltenyi, Auburn, Calif.) to yield DC with >85% purity. DC were then further purified with a fluorescence-activated cell sorter (FACS) (Becton Dickinson and Cytomation, Fort Collins, Colo.) to exclude residual CD3-, CD14-, CD19-, and CD56-staining cells and to separate IPC from MDC. IPC and MDC were subsequently cultured in CM.

HIV strains. HIV-1BaL (from Suzanne Gartner, Mikulas Popovic, and Robert Gallo), HIV-1LAI/IIIB (from Gallo), and human recombinant IL-2 (from Maurice Gately, Hoffmann-La Roche, Inc.) were obtained through the AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health (NIH). HIV-1BaL was prepared by infection of IL-2-stimulated PBMC isolated from buffy coats of HIV-seronegative blood donors. PBMC were infected and then stimulated with 50 U of recombinant IL-2/ml. Virus-containing culture supernatants were harvested 6 days later and stored at -80°C. The 50% tissue culture infective dose (TCID50) was determined in IL-2-stimulated PBMC. HIV-1LAI/IIIB was prepared by infection of T-cell blasts (PBMC stimulated for 3 days with 5 µg of phytohemagglutinin/ml, washed extensively, and resuspended in CM and 50 U of recombinant IL-2/ml). Virus-containing culture supernatants were harvested 6 days later and stored at -80°C. The TCID50 was determined in IL-2-stimulated T-cell blasts.

HIV infection. DC were infected by a 2-h incubation with the macrophage-tropic R5 HIV-1BaL or the T-lymphotropic X4 HIV-1LAI/IIIB strain at 1,500 TCID50s per 106 cells and were then washed three times, resuspended in CM, and plated in 96-well plates at a concentration of 0.2 x 106 cells/ml (40,000 DC/well). When indicated, following the 2-h incubation, soluble CD40L trimer (CD40LT) (gift of Immunex) was added at a final concentration of 500 ng/ml. Cell-free supernatants were collected at 24 h and assessed for IFN-{alpha}, RANTES, and MIP-1{alpha} secretion by enzyme-linked immunosorbent assay (ELISA) (R&D Systems). Cell-free supernatants were collected every 7 days and stored at -80°C for determination of reverse transcriptase (RT) activity and p24 antigen concentration (NEN, Boston, Mass.). Where indicated, IPC were preincubated with RANTES or SDF-1{alpha} at a concentration of 1 µg/ml and were then infected with virus. Where indicated, neutralizing monoclonal anti-IFN-{alpha} antibody (R&D Systems) or a matched isotype control antibody was added to the cultures to a final concentration of 10 µg/ml following infection, and the cultures were supplemented with antibody every 2 days. For viability experiments, infected IPC cultured in parallel with and without CD40LT (40,000 DC/well) were harvested, stained with antibodies, annexin V and PI, and enumerated and analyzed by flow cytometry.

Immunohistochemistry. Following heat-induced epitope retrieval with 10 mM citrate buffer, pH 6.0, formalin-fixed tissue was incubated with anti-human CD123 antibody (Becton Dickinson) and detected with DAKO's (Carpinteria, Calif.) LSAB+ peroxidase system and diaminobenzidine (brown). Double staining was achieved by first staining with anti-CD123 as described but utilizing Vector AS (Vector, Burlingame, Calif.) and then denaturing any residual antibody-enzyme complexes with acid. Following a buffer wash, the tissues were incubated with anti-p24 antibody (DAKO 1:10, overnight at room temperature) and detected with DAKO's LSAB+ Alkaline Phosphatase system with Vector Red (Vector). Paraffin sections from rectal biopsies, cervical biopsies, and tonsil were obtained from the tissue bank at San Francisco General Hospital under an approved human subject protocol. Rectal tissues were derived from routine colonoscopy. Cervical tissues were derived from a patient with cervical dysplasia. The tonsillar tissue was derived from a 30-year-old homosexual male infected with HIV who was on no antiviral medications. He underwent tonsillectomy for adenotonsillar hypertrophy.

IPC/T-cell coculture experiments. Resting CD4+ T cells were purified from autologous PBMC by negative selection against CD8, -14, -19, and -56 and HLA DR with immunomagnetic beads (Miltenyi) and lacked surface expression of CD40L as assessed by flow cytometry. These unstimulated CD4+ T cells were cocultured with HIV-infected DC at a concentration of 0.5 x 106 CD4 T cells/ml (100,000 CD4 T cells/well). In separate cultures, unstimulated CD4+ T cells were also exposed to HIV for 2 h, washed, and cultured without IPC. When indicated, CD40LT was added to the coculture at a final concentration of 500 ng/ml. Cell-free supernatants were collected every 7 days and stored at -80°C for determination of RT activity.

CD4 stimulation by IPC. Naïve CD4 T cells were purified from PBMC of HIV-seronegative study subjects by negative selection for CD8, CD14, CD19, HLA DR, and CD45RO with immunomagnetic beads (Miltenyi). Monocytes and B cells were purified by positive selection with immunomagnetic beads specific for CD14 and CD19, respectively. IPC were purified as previously described. These different antigen-presenting cells were irradiated to 30 Gy and coincubated at 10,000 cells/well with 100,000 autologous CD4+ CD45RA+ T cells/well in CM in 96-well plates. T-cell proliferation in triplicate cultures was assessed after 6 days for [3H]thymidine incorporation over 18 h and measured with a scintillation counter (Wallac, Toruc, Finland).


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RESULTS
 
Circulating DC precursors that include IPC can be identified by their expression of MHC-II and lack of lineage markers and typically represent < 1% of PBMC (Fig. 1a). IPC and MDC precursors can be separated by their expression of CD123 and CD11c, respectively (Fig. 1b). IPC express CD4, CXCR4, and CCR5, requisite coreceptors for HIV, and may therefore be susceptible to HIV infection (Fig. 1c). Moreover, CXCR4 and CCR5 receptors are functional, inducing dose-dependent migration of IPC to their respective ligands, SDF-1{alpha}, and MIP-1{alpha} (Fig. 1d). IPC express high levels of CD62L (Fig. 1c), which is required for migration across high endothelial venules (HEV) (3), and lack expression of DC-SIGN, an HIV binding C-type lectin (12).



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  		FIG. 1. IPC express HIV coreceptors. (a) Blood DC from healthy subjects were phenotypically defined by flow cytometry as expressing MHC-II and lacking lineage markers CD3, -14, -19, and -56 (box). (b) These gated cells were further divided into two distinct subsets either expressing high levels of CD123 and lacking CD11c (IPC) (solid box) or expressing CD11c (MDC) (dashed box). (c) IPC were then further characterized for their surface expression of CD4, CXCR4, CCR5, CD62L, and DC-SIGN. Open histograms represent cells stained with isotype-matched control antibodies. (d) The capacity of IPC (black bar) and MDC (white bar) to migrate in response to SDF-1{alpha} and MIP-1{alpha} was assessed in a transwell chemotaxis assay. Error bars represent the standard deviation for three replicate transwells. (e) Following 10 daily doses of Flt3L to expand DC in vivo, analysis of blood reveals that approximately 16% of PBMC are DC. (f) The proportion of IPC relative to MDC is essentially unchanged by Flt3L treatment. Data shown are representative of three experiments.

Because IPC represent <10% of circulating DC, their isolation in sufficient number for study is challenging. To circumvent the low frequency of IPC in blood, we utilized a growth factor, Flt3 ligand (Flt3L), to expand human DC in vivo (10, 22). In addition to expanding hematopoietic precursors (21), Flt3L may also represent a differentiation factor for IPC (2). Following 10 daily doses of Flt3L, study subjects developed approximately a 10-fold expansion in circulating blood DC (Fig. 1e). The proportion of IPC to MDC remained unchanged following expansion in the total number of DC (Fig. 1f). The pattern of CD4, CXCR4, and CCR5 expression on IPC was not altered by the administration of Flt3L (data not shown).

IPC and MDC were purified from Flt3L-mobilized blood with immunomagnetic beads and separated by FACS on the basis of their expression of CD123 and CD11c to >99% purity. These purified cells represent either IPC (pre-DC2) or immature MDC that lack surface expression of CD80 and 86. We then exposed IPC and MDC to either the macrophage-tropic R5 HIV-1BaL or the T-cell-tropic X4 HIV-1LAI/IIIB strains that use CCR5 or CXCR4, respectively, as a coreceptor for viral entry (1, 11, 30). Macrophage-tropic strains of HIV are responsible for primary HIV infection, while the T-cell-tropic strains usually arise late in infected individuals and are responsible for syncytium formation. After 2 h of incubation, cells were washed free of unbound virus and cultured. HIV replication was assessed by measuring Mg2+-dependent RT activity (19). IPC did not efficiently replicate either HIV-1BaL or HIV-1LAI/IIIB (Fig. 2a). In contrast, MDC that were simultaneously purified replicate HIV-1BaL but not HIV-1LAI/IIIB, consistent with prior reports (23).



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  		FIG.2. HIV replication in IPC requires CD40L activation. IPC and MDC were enriched with immunomagnetic beads and were then separated by FACS based on their differing expression of CD123 and CD11c. IPC and MDC were incubated with HIV for 2 h, washed, and then cultured in parallel in 96 wells at a concentration of 0.2 x 106 cells/ml (40,000 DC/well). (a) Following incubation of IPC and MDC with either HIVLAI/IIIB (triangles) or HIVBaL (circles), triplicate cultures were assessed weekly for viral replication on the basis of RT activity. (b) IPC and MDC were also stimulated with CD40LT following exposure to virus and were assessed weekly for RT activity. (c) IPC exposed or unexposed to HIVBaL and then stimulated with CD40LT were also assessed by flow cytometry for intracellular HIV p24 following 1 week of culture. The histogram on the right represents infected cells stained with an irrelevant isotype-matched control antibody. (d) IPC cultures were also assessed for T-cell contamination and for viability by annexin V and PI staining at day 14. Numbers represent the percentage of cells present in the indicated gates. (e) IPC were incubated with either RANTES or SDF-1{alpha} at a concentration of 1 µg/ml for 1 h prior to and during exposure to medium alone (black bars), HIVBaL (gray bars), or HIVLAI/IIIB (black bars). Cultures were then assessed for p24 production at 1 week by ELISA. Error bars represent standard deviations from triplicate wells. Data shown are representative of three experiments.

We then assessed the capacity of CD40L stimulation to affect HIV replication by activating IPC or MDC with a soluble trimeric CD40L agonist (CD40LT) following exposure of the cells to HIV. CD40/CD40L interactions provide a maturation signal for IPC to differentiate into PDC, increasing the cell surface expression of adhesion, costimulatory, and MHC molecules (7, 15), which are necessary for priming adaptive immunity. Activated CD4 T lymphocytes deliver this signal through cell-cell interaction that would physiologically occur within the T-cell-rich zones of lymphoid tissues (6). IPC were again incubated with the virus for 2 h, washed, and then cultured in medium supplemented with CD40LT. The addition of CD40LT to IPC following viral exposure led to significant replication of both HIV strains (Fig. 2b). MDC infected and then activated with CD40LT also replicated both HIV-1BaL and HIV-1LAI/IIIB, although RT production was significantly reduced relative to MDC exposed to virus without CD40L activation, again consistent with prior reports (23).

To determine what proportion of IPC became productively infected following exposure to virus, we performed intracellular staining for HIV p24 2 weeks after exposure to HIV-1BaL. p24 staining could be detected in over 17% of IPC cultured in the presence of CD40LT (Fig. 2c). At this time point, IPC cultures had minimal contamination by CD3+ T cells (Fig. 2d, upper panel), arguing against contaminating lymphocytes as the source of viral replication. The viability of IPC was not significantly altered by CD40LT or HIV infection (Fig. 2d, lower panel), nor was the number of cells recovered (data not shown). These results argue against CD40LT augmenting viral replication by acting as a survival factor for IPC. HIV-1BaL entry is dependent on CCR5 binding, since preincubation of IPC prior to viral exposure with RANTES/CCL5, a ligand for CCR5, blocked infection while preincubation with SDF-1{alpha}, a ligand for CXCR4, had no effect (Fig. 2e). Conversely, preincubation of IPC with SDF-1{alpha} but not with RANTES blocked infection with HIV-1LAI/IIIB.

To assess the capacity of IPC to produce IFN-{alpha} in response to HIV, we measured IFN-{alpha} in IPC culture supernatants 1 day following addition of HIV-1BaL. As shown in Fig. 3a, the virus-exposed IPC produced significant amounts of IFN-{alpha} that were augmented with CD40L activation. The addition of CD40L without virus also induced IFN-{alpha} secretion by IPC. IPC exposed to supernatants derived from IL-2-stimulated PBMC in the absence of virus did not secrete IFN-{alpha} (data not shown). To determine whether the produced interferon affected viral replication, we cultured HIV-1BaL-infected IPC in the presence or absence of neutralizing anti-IFN-{alpha} antibody. While there was no effect on viral production without CD40LT-induced maturation, addition of anti-IFN-{alpha} antibody significantly accentuated viral replication in IPC incubated with CD40LT (Fig. 3b). IPC were also assessed for their capacity to produce RANTES (Fig. 3c) and MIP-1{alpha}/CCL3 (Fig. 3d) in response to viral infection and/or maturation with CD40LT. While MIP-1{alpha} production was induced only by exposure to HIV, RANTES production could also be induced by CD40LT without virus in a pattern similar to IFN-{alpha} production. Thus, HIV infection leads to activation of IPC that may be distinct from maturation signals delivered through CD40L.



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  		FIG. 3. IFN-{alpha} production by IPC exposed to HIV. (a) IPC production of IFN-{alpha} was measured by ELISA 1 day following exposure to HIVBaL with or without the addition of CD40LT. (b) IPC were also incubated with HIVBaL in the presence or absence of neutralizing anti-IFN-{alpha} antibody, and p24 production was assessed after 1 week. Error bars represent standard deviations from triplicate wells. Data shown are representative of three experiments. IPC production of RANTES (c) and MIP-1{alpha} (d) was measured by ELISA 1 day following exposure to HIVBaL with or without the addition of CD40LT.

We then proceeded to examine the potential role of IPC in HIV pathogenesis in vivo. In addition to blood exposure, HIV can be transmitted via oral, vaginal, and anal exposure. We therefore assessed these sites for the presence of IPC. As has been described, IPC bearing plasmacytoid morphology can be identified in LN based on CD123 staining (8, 15) within the extrafollicular T-cell zones (Fig. 4a). IPC, which express high levels of CD62L, can also be seen in the HEV situated within these zones, consistent with the view that these cells can migrate to LN directly from the circulation. As reported earlier (15), we also found CD123+ cells localized in the parafollicular T-cell-rich zones within the tonsils (Fig. 4b). Significant numbers of IPC were also found in the cervix (Fig. 4c). In rectal biopsies, IPC could be identified in the submucosa but less frequently than in cervix, tonsil, or LN (Fig. 4d).



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  		FIG. 4. Tissue distribution of IPC. (a) IPC can be identified within the extrafollicular T-cell-rich zones of LN by staining with anti-CD123 antibody (brown) (magnification, x100). IPC can be visualized within the HEV. (b) IPC can also be identified in the parafollicular T-cell zone in tonsil (magnification, x100). (c) Large numbers of IPC are seen in the cervical submucosa (magnification, x100). (d) Rare IPC can also be found in the rectal submucosa (arrow) (magnification, x100).

To determine whether HIV is present in IPC in situ, we performed two-color immunohistochemistry (14). Antibody staining without counterstaining was performed on tissue sections to visualize the two colors used: black (anti-CD123) and red (anti-p24). While p24 staining is not seen in uninfected IPC from tonsillar tissue (Fig. 5a), p24-infected cells could be detected in tonsils from an HIV-infected individual (Fig. 5b). Moreover, double-labeled CD123+ p24+ cells could be identified, consistent with HIV infection of IPC in situ (Fig. 5c). The relative frequency of p24+ IPC to that of p24+ CD4 T cells was approximately 50:1. Multinucleated giant cells were also found with both CD123 and p24 staining, consistent with IPC forming syncytia (Fig. 5d).



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  		FIG. 5. Evidence for HIV infection of IPC in vivo. (a) In double-stain experiments without counterstaining of tonsillar tissues, uninfected CD123+ (black) IPC that lack p24 staining (red) can be identified (magnification, x400). (b) p24+ (red) cells can be identified in the tonsils of an HIV-infected individual (magnification, x400). (c) HIV+ IPC can also be seen that are CD123+ (black) and p24+ (red) in tonsil from an HIV-infected individual. (d) Multinucleated giant cells are also identified with CD123 (black) and p24 (red) staining.

The principal targets of HIV are CD4 T cells, which gradually become depleted in infected individuals. To determine whether infected IPC can transmit the virus to these lymphocytes, unstimulated CD4 T cells were cocultured with autologous IPC that had been exposed to HIV and cultured in the absence of CD40LT. While unstimulated CD4 T cells directly exposed to HIV did not replicate virus, CD4 T cells were infected by HIV-exposed IPC, acting in trans (Fig. 6a). Interestingly, replication of HIVBaL was more than twofold greater than that of HIVLAI/IIIB. IPC that had been activated with CD40LT were significantly more efficient than were unstimulated IPC in transmitting virus to CD4 T cells, producing over five times the amount of RT than did IPC/T-cell cocultures without CD40LT stimulation by day 14 (Fig. 6b). Intracellular staining of IPC/T-cell cocultures 1 week following HIVBaL exposure demonstrated that both T cells (CD3+) and IPC (CD3-) produced p24 (data not shown). By comparison, MDC/T-cell cocultures obtained in parallel were up to 50-fold more efficient in replicating virus (Fig. 6c). With MDC, CD40LT accentuated HIVLAI/IIIB transmission but significantly abrogated HIVBaL transmission (Fig. 6d), consistent with a prior report (23). Intracellular staining of IPC/T-cell cocultures 1 week following HIVBaL exposure demonstrated that both T cells (CD3+) and IPC (CD3-) produced p24 (Fig. 6e). Moreover, a subset of IPC in T-cell cocultures lacking CD40LT also was p24+. This finding would be consistent with T cells, which were activated in culture, delivering a CD40L signal to the IPC. The addition of a neutralizing anti-IFN-{alpha} antibody accentuated viral production in IPC/T-cell cocultures, whether or not CD40LT was added (Fig. 6f).



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  		FIG. 6. IPC transmit HIV to autologous CD4 T cells. (a) Following a 2-h exposure to either HIVLAI/IIIB (triangles) or HIVBaL (circles), 4 x 104 IPC were washed and then coincubated with 105 autologous CD4 T cells. Viral replication was assessed weekly for RT activity in triplicate cultures. (b) Infected IPC/T-cell cocultures were also incubated in the presence of CD40LT following exposure to HIV. CD4 T cells that were directly exposed to either HIVLAI/IIIB (squares) or HIVBaL (diamonds), washed, and cultured alone served as controls. Data shown correspond to those for cultures performed in parallel with experiments for Fig. 2 and are representative of three experiments. By comparison, MDC/CD4 T-cell cultures were assessed in parallel either without (c) or with (d) the addition of CD40LT. (e) IPC/T-cell cocultures incubated in the presence or absence of HIVBaL and/or CD40LT were assessed by flow cytometry for intracellular HIV p24 following 1 week of culture. (f) IPC/T-cell cocultures incubated in the presence or absence of neutralizing anti-IFN-{alpha} antibody were assessed for p24 production 1 week following exposure to HIVBaL.

While the immunostimulatory function of MDC has been well characterized, the capacity of IPC to stimulate T cells has only been reported using allogeneic T cells (8, 18, 31). To investigate whether IPC can stimulate naïve autologous CD4 T cells, we cocultured IPC as well as other antigen-presenting cells with naïve T cells in an autologous mixed leukocyte reaction. MDC, IPC, monocytes, and B cells were cultured for 6 days with autologous CD45RA+ CD4+ T cells. IPC, and MDC to a lesser extent, induced proliferation of these naïve CD4 T cells in the absence of exogenous antigen, consistent with an autologous mixed leukocyte reaction (Fig. 7).



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  		FIG. 7. IPC stimulate autologous T cells to proliferate. B cells, IPC, MDC, and monocytes were compared as antigen-presenting cells in an autologous mixed leukocyte reaction. Ten thousand antigen-presenting cells/well were cocultured for 6 days with 100,000 autologous CD45RA+ CD4+ T cells/well. T-cell proliferation was assessed by [3H]thymidine incorporation. Error bars represent standard deviations from triplicate wells. Data shown are representative of three experiments.


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DISCUSSION
 
Relatively little is known about immunopathogenic events occurring during the acute phase of HIV infection. Significant effort has been directed toward determining whether MDC or CD4+ T cells represent the initially infected cells. MDC, including Langerhans cells of the epidermis, have been shown to take up the virus efficiently and infect CD4 T cells in vitro (5, 29). Nevertheless, HIV is inefficiently transmitted through skin exposure. Studies of monkeys exposed orally to simian immunodeficiency virus demonstrated viral replication in CD4+ cells of the tonsil rather than epithelial DC (36); however, CD4+ IPC were not assessed in these experiments. The present study clearly demonstrates that IPC, which are anatomically located at sites of efficient HIV transmission, are susceptible to productive viral infection. These cells express functional chemokine receptors CXCR4 and CCR5, which could potentially recruit IPC to sites of inflammation, such as an infected cervix, and may contribute to the higher efficiency of sexual transmission of HIV in individuals with concurrent sexually transmitted diseases. Moreover, given their expression of CD62L, these infected cells may migrate from the blood directly to lymphoid tissues via HEV. Importantly, in IPC viral replication and the accompanying production of HIV, proteins only occur following maturation with CD40L. This result is in contrast to a prior report (27), in which viral replication was detected in IPC in the absence of CD40L. This report, however, did not address the maturation of the IPC, nor was the purity of IPC discussed. The presence of either CD4 T cells or mature IPC could explain the reported viral replication. IPC maturation induced with tumor necrosis factor alpha also induces viral replication (data not shown) and would be consistent with induction of HIV replication as a consequence of NF-{kappa}B activation. Nevertheless, the requirement of maturation on viral replication differs from the case for MDC, in which viral replication occurs without CD40L-induced maturation and can actually be inhibited by this signal (23). Thus, infected IPC may evade HIV-specific immunity, including cytotoxic T lymphocytes, until they interact with activated CD4 T cells within the extrafollicular T-cell zones of lymphoid tissues. Once this interaction occurs and CD40 is ligated, IPC then replicate virus. IPC may, therefore, serve as viral reservoirs for both macrophage and T-cell-tropic strains.

IPC begin secreting IFN-{alpha} within 24 h of HIV exposure, regardless of whether they replicate virus. Thus, viral infection triggers activation signals distinct from CD40L as has been suggested previously (18). IFN-{alpha} appears to partially suppress viral replication in both IPC and IPC/T-cell cocultures, consistent with prior reports on the effects of IFN-{alpha}/ß on HIV replication (32, 39). While CD40L has been described as a survival factor for MDC, we saw no survival advantage in CD40LT-activated IPC. Cellular viability may have been maintained, in part, by virally induced IFN-{alpha}, which has been shown to be an IPC survival factor (16). Nevertheless, the difference in HIV replication between IPC exposed to HIV in the presence of CD40LT and those exposed in the absence of CD40LT is not due to altered viability of the IPC mediated by CD40 ligation. IPC also produce ß-chemokines in response to viral exposure and therefore would attract CD4 T cells.

Importantly, IPC can infect CD4 T-lymphocytes in trans, even without significant viral replication. Such infection may stem from either internalized or surface-bound virus. CD40L-mediated activation of IPC profoundly increases viral production in the IPC/T-cell cocultures. Since HIV infects proliferating CD4 T cells more efficiently than resting CD4 T cells (13, 37), the ability of IPC to induce naïve T cells to proliferate may also lead to more efficient infection of CD4 T cells. Nevertheless, MDC are approximately 50-fold more efficient than IPC at infecting autologous CD4 T cells. As mentioned previously, the blood-derived IPC used in this study do not express DC-SIGN, which might explain differences in viral replication between IPC and MDC. Further work is necessary to confirm this possibility.

IPC may represent an important HIV reservoir in infected individuals. The capacity of HIV to exploit IPC, a key cell in both innate and adaptive antiviral immunity, could contribute to the chronic and evasive nature of HIV infection. Understanding how CD40 signaling in IPC leads to viral replication should further our understanding of the HIV life cycle and perhaps bring about additional targeted therapies for this devastating viral infection.


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ACKNOWLEDGMENTS
 
We are grateful to Thomas Merigan for making the Stanford Center for AIDS Research available for these studies and to David Parks and the Stanford Shared FACS Facility staff for flow cytometry support.

L.F. is supported by a physician-scientist award from the National Cancer Institute (K23 CA82584-01). M.M. is a fellow of Istituto Superiore di Sanita', Rome, Italy. This investigation was also supported in part by NIH grant P01-HL56443 (to E.G.E.), by Human Health Service grant M01-RR00070 (General Clinical Research Centers, National Center for Research Resources, NIH), by NIH grants MH-59037 and P30 MH59037 UCSF-GIVI CFAR (to B.G.H.), and by NIH grants CA-42509-14 and CA-81543-02 (to L.A. Herzenberg, for support of M.M.).


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FOOTNOTES
 
* Corresponding author. Mailing address: Division of Hematology/Oncology, Department of Medicine, University of California, San Francisco, HSW 513, Box 0511, 513 Parnassus Ave., San Francisco, CA 94143-0511. Phone: (415) 514-3160. Fax: (415) 514-3165. E-mail: lfong{at}medicine.ucsf.edu. Back


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Journal of Virology, November 2002, p. 11033-11041, Vol. 76, No. 21
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.21.11033-11041.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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