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The maturation of dendritic cells (DCs) is associated with a diminished ability to support human immunodeficiency virus (HIV) replication; however, the precise step in the HIV life cycle impaired by DC maturation remains uncertain. Using an HIV virion-based fusion assay, we now show that HIV fusion to monocyte-derived DCs (MDDCs) both decreases and kinetically slows when DCs are induced to mature with poly(I:C) and tumor necrosis factor alpha. Specifically, laboratory-adapted CCR5-tropic 81A virions fused with markedly lower efficiency to mature MDDCs than immature DCs. In contrast, fusion of NL4-3, the isogenic CXCR4-tropic counterpart of 81A, was low in both immature and mature MDDCs. Fusion mediated by primary HIV envelopes, including seven CCR5- and four CXCR4-tropic envelopes, also decreased with DC maturation. The kinetics of virion fusion were also altered by both the state of DC maturation and the coreceptor utilized. Fusion of 81A and NL4-3 virions was delayed in mature compared to immature MDDCs, and NL4-3 fused more slowly than 81A in both mature and immature MDDCs. Surprisingly, primary envelopes with CXCR4 tropism mediated fusion to immature MDDCs with efficiencies similar to those of primary CCR5-tropic envelopes. This result contrasted with the marked preferential fusion of the laboratory-adapted 81A over NL4-3 in immature MDDCs and in ex vivo Langerhans cells, indicating that these laboratory-adapted HIV strains do not fully recapitulate all of the properties of primary HIV isolates. In conclusion, our results demonstrate that the defect in HIV replication observed in mature MDDCs stems at least in part from a decline in viral fusion.
Dendritic cells (DCs) are key cellular players in the pathogenic events associated with human immunodeficiency virus (HIV) infection. DCs localize at sites of viral entry, such as the rectal and vaginal mucosa, and at sites of intense viral replication, such as the lymph nodes (reviewed in reference 38). These cells are thus well positioned to play an important role in regulating the effectiveness of HIV transmission and subsequent viral spread.
The anatomic localization of DCs is intimately linked to their function as antigen-presenting cells. DCs are derived from bone marrow progenitors that home to peripheral mucosal sites, where they differentiate locally into immature DCs. After capturing antigen and under the influence of maturation signals elicited by infection or inflammation, immature DCs undergo a complex cellular maturation process. In vivo, this process is paralleled by DC migration to the lymphoid organs, where the mature DCs efficiently present processed antigenic peptides to interacting T cells (for a review see reference 5).
Increasing evidence suggests that HIV-1 exploits the unique distribution and function of DCs to promote effective viral spread to CD4 T cells. Experiments tracking the entry of simian immunodeficiency virus into the vaginal epithelia of macaque monkeys have suggested that immature epidermal DCs, namely Langerhans cells, are among the first cells to be infected (21, 36). In human vaginal explants exposed to HIV-1, DCs emigrating out of the tissue also carry internalized but intact HIV-1 virions (22). This result suggests that HIV-1 may exploit the natural trafficking properties of DCs for transfer of virions to its primary cellular targets, CD4 T cells, residing in draining lymph nodes. Finally, experiments based on coculture of infected lymphocytes and DCs indicate that conjugates of DCs and T cells form important sites of productive HIV-1 infection (8, 29). Interestingly, the virions produced in such cocultures principally derive from the T cells (15), indicating that DCs can facilitate productive infection of T cells while not themselves serving as hosts for viral replication (2, 18, 25, 27, 28, 42).
DCs can, however, also function as direct cellular targets for HIV infection and can support all phases of the viral life cycle leading to the de novo production of infectious virions (9, 16, 19, 23, 42, 46). Maturation of DCs is associated with a marked decrease in HIV replication; however, the step in the HIV life cycle that is blocked in these mature DCs remains unclear (4, 9, 19). To date, HIV fusion to DCs, one of the earliest steps in this cycle, has only been studied by indirect methods. For fusion to occur, the HIV envelope protein must interact with two cell surface receptors: CD4 and the chemokine receptor CCR5 or CXCR4, which serve as coreceptors for R5- and X4-tropic viruses, respectively (for a review see reference 6). Immature DCs such as Langerhans cells express CD4 molecules and high levels of CCR5 on their surface but not CXCR4, which remains in the intracellular compartments (46). Accordingly, Langerhans cells can efficiently replicate R5-tropic HIV (R5-HIV) in vitro but not X4-tropic strains of HIV (X4-HIV) (23, 46). Interestingly, DC maturation alters the coreceptor expression. Specifically, immature DCs display much higher levels of CCR5 than mature DCs (13, 35). Accordingly, we hypothesized that HIV fusion to DCs might vary with the state of DC maturation. To test this hypothesis, we employed a sensitive and specific flow cytometry-based assay (11, 12) that can detect the fusion of HIV virions to monocyte-derived DCs (MDDCs), primary DCs, and primary CD4 T lymphocytes.
Peripheral blood mononuclear cells (PBMCs) were purified from fresh buffy coats on Ficoll gradients. CD14+ monocytes were positively selected using Miltenyi anti-CD14 magnetic beads according to the manufacturer's instructions. DCs were derived by culturing CD14+ monocytes (2 × 106 cells/ml) for 6 days with 25 ng/ml interleukin-4 (R&D Systems) and 50 ng/ml granulocyte-macrophage colony-stimulating factor (Biosource) in RPMI medium supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, and 100 μg/ml streptomycin (33). The medium was changed every 2 days. DC maturation was induced by incubating the immature DCs for 24 h with 25 μg/ml poly(I:C) (Amersham) and 5 ng/ml tumor necrosis factor alpha (TNF-α) (Biosource) (37). The phenotype of these DCs was verified by immunostaining with antibodies specific for DC (CD1a) and mature DCs (CD40, CD80, CD83, and HLA-DR). Expression of CCR5 and CXCR4 was analyzed by flow cytometry after immunostaining with the 2D7 and the 12G5 antibodies (Becton Dickinson), respectively.
The CCR5 genotype was determined for most of the blood donors. DNA from 105 to 106 cells per donor was purified with the QiaAmp blood kit (QIAGEN). The oligonucleotide primers used to amplify the CCR5 open reading frame products (312 bp for the wild-type allele and 280 bp for Δ32) were GTCTTCATTACACCTGCAGCTCTC (sense) and GGTCCAACCTGTTAGAGCTACTGC (antisense) (32). The PCR amplification was performed in a solution (50 μl) containing 10 to 30 ng of purified genomic DNA, 10 pmol of each primer, 100 μM deoxynucleoside triphosphates, 1 U of Taq DNA polymerase, and 5 μl of PCR buffer containing 10 mM MgCl2 (QIAGEN). The amplification products were analyzed on 2% agarose gels.
Samples of normal skin (10 by 10 cm) were obtained from National Disease Research Interchange. The dermal and epidermal layers were isolated, cut into small pieces, and digested with 7.5 mg/ml collagenases II and IV (Gibco BRL) and 0.3 mg/ml DNase I (Sigma) for 3 h at 37°C. The cell suspension was filtered through a 70-μm mesh and washed twice in RPMI before use.
HIV-1 virions containing the β-lactamase-Vpr (BlaM-Vpr) chimera were produced as previously described (11, 12). Briefly, 293T cells were transfected with pNL4-3 or 81A proviral DNA, pCMV-BlaM-Vpr, and pAdVAntage vectors (Promega). The virions pseudotyped with primary envelopes were produced with pNL4-3Δenv proviral DNA (45) and vectors expressing primary HIV envelope proteins (pSVIII-92RW020.5, pSVIII-92TH014.12, pSVIII-92UG037.8, pSVIII-91US005.11, pSVIII-92BR020.4, pSVIII-92HT593.1, pSVIII-92HT599.24, pSVIII-93MW965.26, pSVIII-92BR025.9, pSVIII-92UG021.16, or pSVIII-92UG024.2, obtained from the National Institutes of Health AIDS Research and Reference Reagent Program). After 48 h of culture at 37°C, the virus-containing supernatant was centrifuged at low speed to remove cellular debris and ultracentrifuged at 72,000 × g for 90 min at 4°C to sediment viral particles. Viral stocks were normalized based on p24Gag content, measured by enzyme-linked immunosorbent assay (NEN Life Science Products).
Slightly different conditions were used to infect the different cell types. MDDCs (2 × 105) and unactivated peripheral blood leukocytes (PBLs) or PBMCs (2 × 106) were infected with HIV (300 to 500 ng of p24Gag) for 1 h at 37°C in 100 μl of RPMI medium. For single-cell suspensions derived from human skin, 106 cells were infected with HIV (2.5 μg of p24Gag). Subsequently, the HIV virion-based fusion assay was performed as previously described (11, 12). Briefly, after incubation of the target cells with virions, the cells were washed once with CO2-independent Dulbecco's modified Eagle medium (DMEM) (Gibco BRL) to remove free virions and loaded with CCF2-AM dye (0.5 mM; Invitrogen) for 1 h at room temperature. After two washes with DMEM, the cells were incubated for 16 h at room temperature in 200 μl of DMEM supplemented with 10% FBS and 2.5 mM probenecid, an inhibitor of anion transport. The cells were next washed once in phosphate-buffered saline (PBS) and fixed in a 1.2% solution of paraformaldehyde overnight. The change in emission fluorescence of CCF2 after cleavage by the BlaM-Vpr chimera was measured by flow cytometry using either a three-laser BD FACSVantage SE instrument or a BD LSRII (Becton Dickinson). Data were collected using FACSDiva software (Becton Dickinson) and analyzed with FlowJo software (Treestar).
NL4-3 or 81A virions containing BlaM-Vpr were first bound to their cellular targets by incubating DCs (107 cells/ml) in 100-μl suspensions of concentrated virions (25 to 50 μg/ml p24Gag) for 1 h at 4°C. After four washes with cold PBS, the cells were resuspended in RPMI with 10% FBS (2.5 × 106 cells/ml) and aliquoted. The aliquots were incubated in tubes in a 37°C water bath for up to 240 min to induce fusion. Fusion was stopped by placing the tubes on ice. The cells were then loaded with CCF2-AM, and fusion was measured as described above. To compare HIV-1 fusion kinetics between donors, we normalized the plateau values obtained (maximal level of fusion) to 100% after subtraction of the background observed at time zero.
When heterogeneous cell populations were examined, the cells to which the virions had fused were phenotyped by immunostaining prior to fixation. Briefly, the cells were washed twice in staining buffer (PBS-2% FBS) and incubated for 30 min at room temperature with the relevant antibodies conjugated to various fluorescent dyes. After two washes, the cells were fixed and analyzed by multiparameter flow cytometry. For phenotyping of T lymphocytes, anti-CD3-allophycocyanin (APC) Cy7 and anti-CD4-phycoeryrthrin (PE) Cy7 antibodies were diluted 1:50 in staining buffer. In coculture experiments, the cocktail included CD1a-APC, CD4-PE Cy7, and CD3-APC Cy7. To identify dermal DCs and Langerhans cells in skin biopsies, the immunostaining cocktail included CD1a-PE, HLA-DR-ECD, CD3-PECy5.5, CD4-PECy7, CD14-APC, and Langerin-APC Cy7. The anti-langerin antibody was conjugated with a Zenon kit according to the manufacturer's instruction (Bioprobes). The compensation was calculated after data collection based on single-stain controls using FlowJo software. To control for immunostaining specificity, we used the fluorescence minus one technique, which corresponds to immunostaining with all of the antibodies except the antibody of interest to determine if increased fluorescence was attributable to staining with the test antibody.
To test associations between the time required to reach 50% of the maximal level of fusion (T50%) and HIV tropism or the state of DC maturation, statistical analyses were performed using the Mann-Whitney U test and Stat-View 5.0 software (SAS Institute).
The HIV virion fusion assay is based on the specific incorporation of a β-lactamase-Vpr (BlaM-Vpr) chimera into HIV virions and its subsequent transfer into the target cell cytoplasm as a result of fusion (11, 12, 39). The BlaM component of the chimera can then enzymatically cleave the CCF2-AM dye loaded into the target cells, changing its fluorescence emission spectrum from green (520 nm) to blue (447 nm). This change in fluorescence emission can be detected by flow cytometry. Importantly, the virion-based fusion assay selectively detects HIV-1 virions entering cells by fusion but not by endocytosis (11).
DCs were differentiated ex vivo from blood monocytes by culturing in the presence of granulocyte-macrophage colony-stimulating factor and interleukin-4 for 6 days. Further DC maturation was induced by treatment with TNF-α and poly(I:C) for 24 h. The phenotype of these different DC populations was verified by immunostaining (Fig. (Fig.1A).1A). As expected, MDDCs expressed high levels of CD1a. Maturation of the DCs was associated with increased expression of CD40, CD80, CD83, and HLA-DR. Such a maturation procedure did not alter the expression of CD4 and CXCR4 but did result in a decline in the expression of CCR5.
First, we compared the susceptibility of immature and mature MDDCs to fusion with a pair of isogenic HIV-1 virions differing in coreceptor tropism, 81A (R5-HIV) and NL4-3 (X4-HIV). As shown in Fig. Fig.1B,1B, fusion of R5-tropic 81A to DCs significantly decreased with DC maturation. For this donor, DC maturation was associated with a slight increase in X4-tropic NL4-3 fusion. As expected, NL4-3 virions fused with higher efficiency to unstimulated CD4 T cells than 81A. Fusion was effectively blocked by the selective inhibitors of TAK779 or AMD3100, a CCR5- or CXCR4-tropic virions, respectively (3) (Fig. (Fig.1C).1C). Similar analysis of DCs from five donors confirmed that DC maturation was significantly associated with diminished fusion of R5-tropic 81A (P = 0.0079), while no significant changes were observed in X4-tropic NL4-3 fusion (Fig. (Fig.1D).1D). Maturation also decreased fusion of YU-2, another laboratory-adapted R5-tropic HIV strain (data not shown). HIV-1 binding to alternative receptors like C-type lectin receptors might influence the levels of fusion. To explore this possibility, we tested the effects of mannan, a C-type lectin receptor ligand that decreases HIV-1 binding to immature DCs (41, 42) and inhibits HIV-1 replication (23). Fusion of 81A virions to immature MDDCs was sharply decreased in the presence of mannan; less pronounced effects were observed for NL4-3 virion fusion to immature MDDCs and for fusion of both types of virions to mature MDDCs (Fig. (Fig.1E).1E). Mannan had no effect on fusion to CD4 T lymphocytes (data not shown). Thus, C-type lectin receptors appear to play a role in the singularly efficient fusion of R5-tropic HIV-1 to immature MDDCs.
We next evaluated whether 81A fuses efficiently to primary DCs, specifically epidermal Langerhans cells and dermal DCs. These two cell types were identified in single-cell suspensions prepared from skin biopsy samples utilizing a six-color immunostaining strategy. Both cell types express high levels of HLA-DR and CD1a, intermediate levels of CD4, and no CD3 or CD14. The Langerhans cells uniquely express langerin. Fluorescence minus one controls were used to monitor the specificity of immunostaining (Fig. (Fig.2A).2A). Like the immature MDDCs, both Langerhans and dermal DCs supported much higher levels of fusion of R5-tropic 81A than X4-tropic NL4-3 (Fig. (Fig.2B).2B). Similar results were obtained with single-cell suspensions prepared from skin samples from three different donors (Fig. (Fig.2C).2C). Thus, R5-tropic 81A virions fuse to immature DCs, including primary Langerhans cells and dermal DCs, with far greater efficiency than X4-tropic NL4-3 virions.
Next, we assessed the effects of DC maturation and HIV-1 coreceptor tropism on the kinetics of virion fusion, measured as the time required for cell-surface-bound virions to recruit HIV-1 receptors and coreceptors and successfully fuse to target cells. Virions were initially bound to cells at 4°C and then shifted to 37°C for various times. Maximal levels of HIV fusion with each viral preparation and cell type are shown in Fig. Fig.3A.3A. As expected, no fusion events were detected in cells held at 4°C (data not shown). Measurements of fusion over time revealed a marked effect of both the state of DC maturation and HIV-1 coreceptor tropism on the kinetics of virion fusion (Fig. (Fig.3B).3B). Specifically, both R5- and X4-tropic HIV virions fused to immature MDDCs more rapidly than to mature MDDCs, and 81A virions fused more rapidly than NL4-3 virions. In contrast, 81A and NL4-3 virions fused to CD4 T lymphocytes with indistinguishable kinetics (Fig. 3C and D). Similar results were obtained with cells isolated from six independent donors (Fig. (Fig.3E).3E). As determined by the time required to reach 50% of the maximal level of fusion (T50%), HIV fused significantly faster to immature DCs than to mature DCs (P < 0.005).
The particularly rapid kinetics of fusion of R5-tropic 81A to immature MDDCs could reflect the high surface levels of CCR5 present on these cells. Alternatively, this finding could result from a more efficient incorporation of Env or BlaM-Vpr into 81A virions or from the higher absolute levels of fusion observed with the immature MDDCs. Immunoblotting studies revealed that 81A virions did not incorporate more gp41 or BlaM-Vpr than NL4-3 virions (Fig. (Fig.4A),4A), and the rapid fusion of 81A to immature DCs persisted even when the viral preparation was serially diluted to reduce the absolute level of fusion attained (Fig. 4B and C). Finally, we examined whether the high level of CCR5 was responsible for the fast kinetics of fusion by titrating these receptors with the CCR5 antagonist, TAK-779 (Fig. 4D and E). As expected, TAK-779 decreased the number of cells supporting fusion in a dose-dependent manner (Fig. (Fig.4D).4D). Moreover, the kinetics of fusion were progressively slowed as increasing doses of this inhibitor were added (Fig. (Fig.4E).4E). Thus, the kinetics of HIV-1 fusion in MDDCs appear to be dictated by both the state of DC maturation and the tropism of HIV-1. The fast fusion kinetics observed with R5-tropic 81A and immature MDDCs also relates at least in part to the high levels of CCR5 expressed on these cells.
Finally, we investigated whether DC maturation also affects fusion mediated by envelopes from primary HIV isolates. A panel of 11 primary envelopes was obtained from the AIDS Research and Reference Reagent Program (17) and used to pseudotype HIV virions (Table (Table1).1). First, we verified the tropism of these envelopes using freshly isolated PBLs and AMD3100 or TAK-779 (Fig. (Fig.5A).5A). All of the envelopes displayed the expected tropism, except the dual-tropic 92TH593.1, which was mainly sensitive to AMD3100 and insensitive to TAK-779. Next, we compared the ability of these envelopes to mediate fusion to immature and mature MDDCs. Maturation significantly decreased fusion mediated by all these envelopes (P < 0.005). However, in contrast to results obtained with the laboratory-adapted viruses, primary envelopes displaying X4 tropism mediated fusion to immature MDDCs as efficiently as R5-tropic envelopes.
Our studies demonstrate that the maturation of MDDCs both decreases the magnitude and kinetically slows the rate of HIV virion fusion. Specifically, laboratory-adapted 81A virions and virions pseudotyped with various primary envelopes fused with lower efficiency to mature than immature MDDCs, and the fusion kinetics of 81A and NL4-3 were slower in mature MDDCs than in immature MDDCs.
These findings confirm and extend the results of Granelli-Piperno et al., who showed by quantification of the early products of reverse transcription that maturation of MDDCs decreased the entry of the R5-tropic Ba-L (19). However, our results contrast with the increased entry of Ba-L and the X4-tropic LAI observed in mature DCs derived in vitro from CD34+ hematopoietic progenitor cells (9) and with the changes in Ba-L and LAI entry in MDDCs (4). The origin of the viruses used in our studies cannot easily account for these differences, since 81A contains the V1 to V3 loop of Ba-L (40) and LAI and NL4-3 encode identical envelopes (1). These contradictory results could reflect the nature of the DCs and differences in procedures for inducing DC maturation and measuring viral fusion. In our experiments, immature MDDC resembled primary Langerhans cells and dermal DCs in their sensitivity to 81A and NL4-3 fusion, indicating that MDDCs function as an acceptable surrogate for DCs present in tissues in vivo. Furthermore, treatment of MDDCs with poly(I:C) and TNF-α induced complete phenotypic maturation, avoiding the generation of a heterogenous population which could potentially mask the effect of maturation on HIV fusion. Finally, our measurement of viral fusion is not affected by the abundance of virion endocytosis that characterizes DCs, since the HIV virion-based fusion assay, but not the use of early products of reverse transcription as an “entry marker”, measures viral fusion independently of viral endocytosis (11). Therefore, our results strongly suggest that in vivo fusion of HIV to mature DCs is also impaired.
HIV-1 binding to alternative receptors like C-type lectin receptors influences HIV-1 replication in DCs (26, 41, 42). We found that mannan, one of the ligands of the C-type lectin receptor, significantly decreased fusion of R5-tropic virions to immature MDDC; the effect was not as pronounced for the X4-tropic NL4-3 virions. Mannan had no effect on HIV-1 fusion to mature MDDCs, consistent with the lower amount of C-type lectin receptors on mature DCs (14). Thus, C-type lectin receptors appear to mainly play a role in the efficient fusion of R5-tropic HIV-1 to immature MDDCs. Previous studies based on immunohistochemical and confocal microscopy had indicated that DC-SIGN, one of the C-type lectin receptors, colocalizes with CD4 and CCR5 on alveolar macrophages (26), suggesting that C-type lectin receptors could focus HIV-1 virions in a location that favors successful engagement of CD4 and CCR5 and not CXCR4. This location could be the lipid rafts, since DC-SIGN, CD4, and CCR5 are preferentially enriched in lipid rafts compared to CXCR4 (7, 30).
In addition to the changes in the overall level of HIV fusion, laboratory-adapted strains of HIV also fused more rapidly to immature MDDCs than to mature MDDCs. The rapid fusion kinetics displayed by R5-tropic 81A virions relates in part to the high density of CCR5 receptors at the plasma membrane, since slower kinetics were observed when available CCR5 receptors were reduced by the addition of graded doses of the CCR5 antagonist TAK-779. Maturation of MDDCs, which is intrinsically associated with a decline in CCR5 expression (13, 35), also led to lower levels and slower kinetics of R5-tropic 81A fusion. Interestingly, although DC maturation induced by poly(I:C) and TNF-α did not decrease cell surface expression of CXCR4 or CD4, the kinetics of fusion of the X4-tropic NL4-3 to mature MDDCs was also delayed in these cells. Morphological changes induced by DC maturation could be responsible for this finding. Specifically, the development of cellular dendrites in mature MDDCs (10) could separate microclusters of CD4, usually present at the tip of the dendrite, from the coreceptors, which usually localize near the base of the dendrite (34, 44). Consequently, gp120/gp41 complexes bound to CD4 might require more time to effectively engage the coreceptors and trigger the fusion reaction.
Surprisingly and in sharp contrast to the laboratory-adapted viruses, primary envelopes with CXCR4 tropism mediated fusion to immature MDDCs with efficiencies similar to those of R5-tropic primary envelopes. These results contrasted with the ~20- to 70-fold difference in fusion observed in immature MDDCs with the R5-tropic 81A and X4-tropic NL4-3 laboratory viruses. Despite lower levels of fusion, the four X4-tropic primary envelopes and the seven R5-tropic envelopes mediated comparable levels of fusion in immature MDDCs obtained from multiple donors. It is very unlikely that the pseudotyping procedure differentially altered the properties of R5- and X4-tropic envelopes. Therefore, these surprising results suggest that primary viruses with tropism for either CCR5 or CXCR4 may fuse similarly in vivo to immature DCs, such as Langerhans cells. Previous studies of HIV replication in Langerhans cells relied on the use of laboratory-adapted strains of HIV and showed high replication of R5-HIV and not X4-HIV, suggesting that immature DCs might play a role in the preferential transmission of R5-HIV (20, 23, 24, 31). However, another study noted that immature MDDCs exposed to HIV-1 isolates with mixed tropism did not exclusively replicate the R5-tropic isolates (43). Although our results with laboratory-adapted strains of HIV support a role of immature DCs in the preferential transmission of R5-HIV, our more physiologically relevant analysis of fusion with primary envelopes does not support this model.
In summary, we have shown that the maturation of dendritic cells is associated with a marked decline and slowing of HIV fusion. Since DC maturation also alters HIV transcription (4), the fusion defect could be the first of several blocks encountered by viruses in these cells that culminate in reduced viral replication in mature DCs. This defect in fusion, which leaves HIV virions intact, could potentially facilitate handling of the virus by endocytosis with later transfer of these intact virions to interacting CD4 T cells.
We thank Dominique Schols for the gift of AMD3100, Stuart Turville, Wes Yonemoto, Tim Beaumont, and David Favre for stimulating discussions, Marty Bigos for assistance in the flow cytometry-based experiments, Stephen Ordway and Gary Howard for editorial assistance, John C. W. Carroll for graphic arts, and Sue Cammack and Robin Givens for administrative assistance. We also thank the donors of blood and skin biopsies and the NIH AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH, for the vectors expressing various primary envelops and the monoclonal antibody to gp41.
We also gratefully acknowledge funding support for these studies from the National Institutes of Health (P01 HD40543 and R03 AI062263-01A) and the University-Wide AIDS Research Program (C99-SF-02 and F03-GI-205).