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Journal of Virology, May 2005, p. 5762-5773, Vol. 79, No. 9
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.9.5762-5773.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Binding and Transfer of Human Immunodeficiency Virus by DC-SIGN⁺ Cells in Human Rectal Mucosa

Kevin B. Gurney,¹ Julie Elliott,^2,3 Hoorig Nassanian,¹ Carol Song,¹ Elizabeth Soilleux,⁴ Ian McGowan,^2,3 Peter A. Anton,^2,3* and Benhur Lee^1,3,5*

Department of Microbiology, Immunology and Molecular Genetics,¹ Department of Medicine,² UCLA AIDS Institute,³ Department of Pathology and Laboratory Medicine, David Geffen School of Medicine, UCLA, Los Angeles, California 90095-1489,⁵ Medical Research Council Cancer Cell Unit, Hutchison/MRC Research Centre, Cambridge CB2 2XY, United Kingdom⁴

Received 30 June 2004/ Accepted 21 November 2004

Top Abstract Introduction Materials and Methods Results Discussion References

 
The role of DC-SIGN on human rectal mucosal dendritic cells is unknown. Using highly purified human rectal mucosal DC-SIGN⁺ cells and an ultrasensitive real-time reverse transcription-PCR assay to quantify virus binding, we found that HLA-DR⁺/DC-SIGN⁺ cells can bind and transfer more virus than the HLA-DR⁺/DC-SIGN^– cells. Greater than 90% of the virus bound to total mucosal mononuclear cells (MMCs) was accounted for by the DC-SIGN⁺ cells, which comprise only 1 to 5% of total MMCs. Significantly, anti-DC-SIGN antibodies blocked 90% of the virus binding when more-physiologic amounts of virus inoculum were used. DC-SIGN expression in the rectal mucosa was significantly correlated with the interleukin-10 (IL-10)/IL-12 ratio (r = 0.58, P < 0.002; n = 26) among human immunodeficiency virus (HIV)-positive patients. Ex vivo and in vitro data implicate the role of IL-10 in upregulating DC-SIGN expression and downregulating expression of the costimulatory molecules CD80/CD86. Dendritic cells derived from monocytes (MDDCs) in the presence of IL-10 render the MDDCs less responsive to maturation stimuli, such as lipopolysaccharide and tumor necrosis factor alpha, and migration to the CCR7 ligand macrophage inflammatory protein 3ß. Thus, an increased IL-10 environment could render DC-SIGN⁺ cells less immunostimulatory and migratory, thereby dampening an effective immune response. DC-SIGN and the IL-10/IL-12 axis may play significant roles in the mucosal transmission and pathogenesis of HIV type 1.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Dendritic cells (DCs) are considered sentinels of the immune system, positioned at peripheral sites and poised to capture pathogens immediately upon exposure (2). Under the inflammatory conditions that arise when pathogens gain access to the host, antigen-laden DCs are signaled to traffic back to the draining lymph node to process and present the foreign antigens to the diverse pool of naïve T cells. Pathogen-specific helper T cells and cytotoxic T cells are specifically stimulated, thereby initiating the adaptive immune response (37). It has been proposed that human immunodeficiency virus (HIV) takes advantage of the trafficking pattern of DCs to gain access to the lymph nodes for viral propagation and dissemination (17, 47, 54). A subset of DCs present in dermal and mucosal tissues express the C-type (calcium-dependent) lectin, DC-specific ICAM-3-grabbing nonintegrin (DC-SIGN), which has been shown to bind to the mannose residues on the heavily glycosylated gp120 protein (21). Other lectin receptors reportedly have HIV binding affinity, such as mannose receptor and langerin (53), which are expressed on intradermal DCs and Langerhans cells, respectively. However, since DC-SIGN on DCs has been shown to bind and transfer HIV to permissive T cells (10, 17), much attention has been paid to the DC-SIGN interaction with gp120, as disruption of this interaction represents a potential new therapeutic approach for preventing HIV transmission.

Since DCs are a rare population in the peripheral tissues and are thus difficult to isolate, many have turned to the more tractable model of in vitro monocyte-derived DCs (MDDCs), which express abundant levels of DC-SIGN, for study. Yet, characterization of HIV interaction with the primary mucosal DCs implicated in the transmission process is lacking. To address this need, we acquired primary gut mucosal DC-SIGN⁺ cells from human rectal mucosal biopsy tissue for functional and phenotypic characterization.

The gut mucosal tissue is also the largest repository of immune cells and is a highly permissive environment for HIV replication (32, 55). This replication persists even in the presence of highly active antiretroviral therapy that suppresses viral replication in the peripheral blood (1, 6, 28, 29). This may be due in part to the greater activation state of gut-associated lymphocytes compared to those in peripheral blood and the spleen, due to constant exposure to microbial and dietary antigens (41). Such an environment necessitates the existence of mechanisms to exert a greater tolerogenic potential in gut immune cells in order to prevent chronic activation. Indeed, murine colonic DCs express greater levels of the regulatory cytokine interleukin-10 (IL-10) than DCs from the spleen and blood (22). A break from this tolerogenic state to an activated Th-1-like inflammatory state is associated with inflammatory bowel diseases (40). Thus, maintaining this tolerogenic state to prevent inflammation and immune activation is an attractive target for pathogen subsistence, as is the case for chronic infections (31). Interestingly, other pathogen-derived ligands to DC-SIGN, such as the lipoarabinomannan component of the Mycobacterium tuberculosis cell wall, have been shown to trigger DC IL-10 secretion via specific interactions with DC-SIGN (18). Thus, we also sought to characterize the immunological environment that might modulate DC-SIGN expression during established HIV infection in the gut. As an immune-regulatory cytokine, IL-10 has been shown to decrease costimulatory molecule expression on DCs and impair DC maturation and migration (7, 11). Here, we provide data that suggest a role for the regulatory cytokine IL-10 in inducing an immunosuppressive environment in vivo and further show the unique ability of IL-10 to induce high levels of DC-SIGN surface expression in vitro in MDDCs. Thus, DC-SIGN and the IL-10/IL-12 axis may have biological relevance in the mucosal transmission and pathogenesis of HIV type 1 (HIV-1).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Antibodies and reagents. HLA-DR-fluorescein isothiocyanate (FITC) and HLA-DR-Tri-Color (TC), CD19-allophycocyanin (APC), CD3-APC, CD45-TC, CD14-APC, CD11c-APC, CD40-phycoerythrin (PE), CD80-PE, and CD86-PE antibodies were obtained from Caltag (Burlingame, CA). BDCA-3-PE, BDCA-4-PE, and BDCA-4-APC antibodies were obtained from Miltenyi Biotec (Auburn, CA). CD3-FITC antibody was obtained from Beckman Coulter (Miami, FL). CD56-APC, macrophage mannose receptor-PE, CCR6-PE, CD45-PerCP, CCR5-PE, and CXCR4-PE were obtained from BD-Pharmingen (San Jose, CA). DC-SIGN-PE antibody was obtained from R&D Systems (Minneapolis, MN). Antibody to the repeat region of DC-SIGN (DC-28) was labeled with either Zenon-Alexa488 or Zenon-PE from Molecular Probes (Eugene, OR) for immunophenotyping and/or fluorescence-activated cell sorting so as to not hinder virus binding to the C-terminal carbohydrate recognition domain of DC-SIGN. The cytokines IL-4, IL-10, IL-13, macrophage inflammatory protein 3ß (MIP-3ß), tumor necrosis factor alpha (TNF-), and granulocyte-macrophage colony-stimulating factor (GM-CSF) were obtained from Peprotech (Rocky Hill, NJ). PHA-P and bacterial lipopolysaccharide (LPS) were obtained from Sigma (St. Louis, MO). Recombinant human IL-2 was obtained through the AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health (NIH). Human recombinant IL-2 was obtained from Maurice Gately, Hoffmann-La Roche Inc. (26).

Cells. Monocytes (for in vitro differentiation to DCs) and CD4 T cells were isolated from the peripheral blood of normal healthy donors. Both monocytes and CD4 T cells were isolated by using RosetteSep (StemCell, Vancouver, BC) according to the manufacturer's guidelines. Monocytes were diluted to a concentration of 0.8 to 1.6 million cells/ml in RPMI medium (Life Technologies) containing 10% fetal bovine serum (SeraCare, Oceanside, CA) and penicillin/streptomycin (Life Technologies/Invitrogen, Carlsbad CA) supplemented with IL-4 (100 ng/ml) and GM-CSF (50 ng/ml) and plated in 24- or 12-well plates at a 250-µl and 500-µl volume, respectively. MDDC maturation was induced with LPS (10 ng/ml) and TNF- (100 ng/ml). Primary rectal mucosal mononuclear cells (MMCs) were obtained via flexible sigmoidoscopy from 30 cm in the rectosigmoid colon from otherwise-healthy, stable HIV⁺ and healthy HIV^– patients without gastrointestinal diseases according to institutional review board guidelines and informed consent, as previously described (43).

Virus preparation. The replication-competent R5 HIV-1, JR-CSF, was prepared by transfecting the plasmid pYK-JRCSF into HEK 293T cells. Pseudotyped HIV-1 (green fluorescent protein [GFP] reporter) with SIV316 Env was performed by cotransfection of 293T cells with pNL-GFP and a plasmid containing SIV316 Env at a 1:3 ratio of plasmids. Forty-eight hours posttransfection, the viral supernatants were collected and filtered through 0.22-µm filters and frozen at –80°C. The viral p24 level in the supernatant was determined as a measure of virus titer.

Cell sorting. For viral binding studies, total mucosal mononuclear cells were labeled with HLA-DR-APC and DC-SIGN-PE (Zenon anti-immunoglobulin G2a [IgG2a] PE-labeled DC28 antibody to the repeat region of DC-SIGN) and then sorted for HLA-DR⁺/DC-SIGN⁺ and HLA-DR⁺/DC-SIGN^– cells by using a fluorescence-activated cell sorter (FACS) Vantage SE apparatus with the FACS DiVa option (Becton-Dickinson, San Jose, CA). For virus transfer experiments, MMCs were additionally labeled with CD3-FITC and sorted for the same populations, but the CD3⁺ T cells were excluded. This was to ensure that the viral transfer only occurred to the allogeneic CD4⁺ T-cell blasts that were added to the sorted DC-SIGN⁺ and DC-SIGN^– cells.

Virus binding assay. Prior to incubating the sorted mononuclear populations with virus, the cells were preincubated with or without one of the following blocking agents: mannan (Sigma, St. Louis, MO) at 5 mg/ml or anti-DC-SIGN antibody (clone 612; R&D Systems, Minneapolis, MN) at 10 µg/ml for 30 min at 4°C. Virus at 70 to 100 ng of p24 was added per 100,000 sorted cells and incubated for 2 h at 37°C, and the cells were then washed four times with medium to remove unbound virus. Each sample was frozen at –80°C in RNA lysing buffer (Stratagene, La Jolla, CA). RNA was isolated, and the number of virions bound per cell was determined by performing quantitative real-time reverse transcription-PCR (RT-PCR) for viral genomic RNA (see "Quantitative RT-PCR," below).

Virus binding to the total population was performed with less virus than with the sorted population. HIV-1 (JR-CSF) at 0.5 to 12.5 ng of p24 was added per 100,000 gut mucosal mononuclear cells and incubated for 2 h. Excess virus was removed by washing the cells four times with medium, and the total MMCs were stained for HLA-DR and DC-SIGN. The HLA-DR⁺/DC-SIGN⁺ and HLA-DR⁺/DC-SIGN^– cells were sorted and then frozen in RNA lysing buffer. MMCs not passed through the sorter were also collected and frozen in RNA lysing buffer for quantitative RT-PCR analysis.

Virus was titrated on MDDCs to determine the sensitivity of the RT-PCR assay. HIV-1 (JR-CSF) was added at the ranges of 500 pg, 50 pg, and 5 pg of p24 to 50,000 MDDCs for 2 h at 37°C to cells preincubated for 30 min at 4°C with the following inhibitors: mannan (Sigma, St Louis, MO) at 5 mg/ml, anti-DC-SIGN antibody (clone 612; R&D Systems, Minneapolis, MN) at 10 µg/ml, anti-CD4 at 10 µg/ml (clone RPA-T4; BD-Pharmingen, La Jolla, CA), or mouse IgG1 at 10 µg/ml (Beckman Coulter, Miami, FL). After 2 h of incubation, cells were washed three to four times and the cells were lysed for RNA isolation.

RNA isolation. The Nanoprep RNA isolation kit (Stratagene, La Jolla, CA) was used to isolate RNA from the small number of FACS-sorted DC-SIGN⁺ and DC-SIGN^– cells. Contaminating DNA was digested on the Nanoprep columns according to the manufacturer's guidelines. For cytokine RT-PCR, RNA was extracted from gut mucosal tissue sections using a modification of the TRIzol isolation protocol (Invitrogen, Carlsbad, CA). Tissue biopsies were homogenized in 1 ml of TRIzol using a Powergen homogenizer (Fisher Scientific, Pittsburgh, PA) fitted with sterile disposable generators. The aqueous phase was collected following centrifugation and placed on an RNeasy column (QIAGEN, Valencia, CA) for further isolation. Finally, the RNA was eluted with RNase-free water and treated with DNA-free (Ambion, Austin, TX) to remove any contaminating DNA.

Quantitative RT-PCR. Quantitative real-time RT-PCR was performed on the isolated RNA by using the Quantitect probe RT-PCR kit (QIAGEN, Valencia, CA) on the DNA Engine Opticon Monitor 2 (MJ Research Inc, South San Francisco, CA). For HIV genomic RNA detection, we used the long terminal repeat (LTR) forward primer (5'-AACTAGGGAACCCACTGCTTAAG-3'), LTR reverse primer (5'-CTCCTAGAGATTTTCCACACTGACTAA-3'), and the fluorogenic probe (6-carboxyfluorescein [6FAM]-5'-TTACCAGAGTCACACAACAGACGGGCA-3'-tetramethyl carboxyrhodamine [TAMRA]) in the RT-PCR. The quantity of HIV was calculated by interpolation from a standard curve generated by running in parallel serial dilutions of known quantities of the HIV plasmid pYKJR-CSF. The HIV signals were normalized against the housekeeping gene ß-actin using the ß-actin forward primer (5'-GCATGGGTCAGAAGGATTCCT-3'), ß-actin reverse primer (5'-TGCCAGATTTTCTCCATGTC-3'), and the fluorogenic probe (6FAM-5'-TGAAGTACCCCATCGAGCACGGCAT-3'-TAMRA). The ß-actin copy number was also calculated by interpolation from a standard curve generated from serial dilutions of a plasmid containing ß-actin cDNA (IMAGE clone 2900526; Invitrogen, Carlsbad, CA).

Cytokine mRNA quantification was performed in a two-step RT-PCR protocol. Total RNA was reverse transcribed into cDNA using random primers according to the ProSTAR first-strand RT-PCR kit protocol (Stratagene, La Jolla, CA) and then amplified using Amplitaq Gold DNA polymerase according to the universal PCR Taqman mix conditions (Applied Biosystems, Foster City, CA) on the GeneAmp 5700 sequence detection system (Applied Biosystems, Foster City, CA). IL-12 p40 message was amplified and detected by the forward primer (5'-ACCCAACAACTTGCAGCTGAA-3'), reverse primer (5'-TGGACCTGAACGCAGAATGTC-3'), and fluorogenic probe (6FAM-5'-TCAGCTGGGAGTACCCTGACACCT-3'-TAMRA). IL-10 message was amplified and detected by the forward primer (5'-GCTGAGAACCAAGACCCAGAC-3'), the reverse primer (5'-GGAAGAAATCGATGACAGCG-3'), and the fluorogenic probe (6FAM-5'-CCCTGTGAAAACAAGAGCAAGGCCG-3'-TAMRA).

Virus transfer assay. Sorted DC-SIGN⁺ and DC-SIGN^– cells from the HLA-DR⁺/CD3^– gate were added to a 96-well plate at 16,000 cells per well. Twice the number of CD4⁺ T cells stimulated prior for 2 days in IL-2 (1,000 IU/ml) and PHA-P (5 µg/ml) were added to the cell cultures for a combined volume of 150 µl. The supernatant was sampled at days 1, 4, and 7 to measure viral p24 levels by enzyme-linked immunosorbent assay (Coulter, Miami, FL).

Chemotaxis assay. Six hundred microliters of RPMI medium containing 10% fetal bovine serum with or with out MIP-3ß at 250 ng/ml was placed in the bottom well of a 24-well transwell plate (Costar, Corning, NY). One hundred microliters of MDDCs at 6 x 10⁵ to 10 x 10⁵ cells/ml was placed in the top insert with a pore size of 5.0 µm. Migration took place at 37°C for 3 to 4 h, after which 500 µl of the bottom well was collected and the number of cells that passed through was counted on a flow cytometer. The amount of chemotaxis to the MIP-3ß gradient was expressed as a percent relative to migration that occurred in the absence of MIP-3ß under each MDDC condition.

Immunofluorescence. Formalin-fixed tissue from gut mucosal biopsies were cut in 5-µm sections and subjected to an antigen retrieval process as described previously (45). A primary rabbit antibody to the C terminus of DC-SIGN was used to stain for DC-SIGN. The rabbit antibodies were detected with a goat-anti-rabbit secondary antibody conjugated to Alexa Fluor 594 (Molecular Probes, Eugene OR). Dual staining for DC-SIGN and CD14 was performed with a mouse anti-human DC-SIGN (clone 28) followed by goat anti-mouse Alexa Fluor 488 (Molecular Probes, Eugene, OR) and sheep anti-human CD14 (R&D Systems, Minneapolis, MN) followed by donkey anti-sheep Alexa Fluor 594 (Molecular Probes, Eugene, OR). Fluorescent images were captured using a Nikon Eclipse TE300 microscope (Melville, NY), and the number of DC-SIGN⁺ cells was enumerated by the Metamorph imaging analysis software (Universal Imaging Corporation, Downington, PA). The operator was blinded as to the HIV status of the patient sample, and the total morphometric analysis was performed by two independent operators.

Statistical analysis. Comparisons of DC-SIGN counts between HIV⁺ and HIV^– samples were performed using Student's t test (two-tailed, two-sample unequal variance). The Pearson's correlation (r), the P value of the correlation, and the 95% confidence interval of the correlation boundary were calculated using the GraphPad Prism software (San Diego, CA). The Bonferroni inequality was used to confirm that the P value for the correlative studies remained significant (<0.05) when performing multiple correlations between the various cytokines and DC-SIGN counts. For the number of comparisons used (five), a P value of <0.01 was used as a threshold for significance.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Phenotype of DC-SIGN⁺ cells in the gut. DC-SIGN-expressing cells are found in the lamina propria and subepithelial dome of Peyer's patches of the human gut mucosa (23, 45), and it has been posited that DC-SIGN⁺ DCs in the submucosa serve as a conduit for the transfer of HIV-1 from the periphery to the draining lymph nodes during primary sexual mucosal transmission of HIV-1 (17, 46). Approximately 1 to 5% of the total rectal MMCs are DC-SIGN⁺ cells, whereas in the blood, DC-SIGN⁺ cells make up less than 0.01% of peripheral blood mononuclear cells (14) (Fig. 1). Rectal mucosal DC-SIGN⁺ cells expressed the highest intensity of major histocompatibility complex class II (HLA-DR) of the total MMC population and were found exclusively in the CD45⁺ hematopoietic-derived population (Fig. 1A). DC-SIGN⁺ mucosal cells expressed the costimulatory molecules CD80, CD86, and CD40. Like tissue macrophages, DC-SIGN⁺ cells expressed CD14, the macrophage mannose receptor, and CD11c. The coexpression of CD14 on a majority of DC-SIGN⁺ cells in the gut was confirmed by immunofluorescence (Fig. 1C). DC-SIGN⁺ cells also expressed the CCR6 chemokine receptor, characteristic of peripheral tissue-homing DCs (12). As for HIV-1 entry receptors, DC-SIGN⁺ cells expressed CD4 but undetectable levels of the chemokine receptors CCR5 and CXCR4, despite clear expression of CCR5 and CXCR4 on the gut mucosal lineage-positive cells from the same sample (47% and 60%, respectively) (Fig. 1B). The DC-SIGN⁺ cells appeared to be immature DCs, in that they did not express the DC maturation molecule CD83. Furthermore, DC-SIGN⁺ cells did not express the high levels of CD123 seen on CD123^high, HLA-DR⁺ plasmacytoid DCs in the blood, despite the presence of BDCA-4 (Fig. 1B), which is also a marker for plasmacytoid DCs in the blood (13). However, unlike BDCA-2, BDCA-4 is also expressed on MDDCs and, thus, is not considered a unique marker for plasmacytoid DCs (13). Furthermore, the other pDC marker, BDCA-2, was not expressed on DC-SIGN⁺ cells in the gut (Fig. 1B).

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FIG. 1. Phenotype of DC-SIGN+ cells in the gut mucosa. (A) The rectal MMC suspension was stained with HLA-DR-FITC, IgG2b-PE, and CD45-PerCP or HLA-DR-FITC, DC-SIGN-PE, and CD45-PerCP. A gate was drawn on the HLA-DR+/DC-SIGN+ cells based on the isotype control. Total mucosal cells were displayed based on CD45 and side scatter (SSC), with another gate drawn on where the HLA-DR+/DC-SIGN+ cells lie to show its large granularity and association with the hematopoietic cell (CD45) population (arrows). (B) MMCs from three HIV+ and three HIV– individuals were stained with a lineage cocktail (CD19-APC, CD3-APC, and CD56-APC), HLA-DR-TC, DC-SIGN antibody (clone 28) conjugated with Alexa Fluor 488 using the Zenon mouse IgG2a labeling kit, and the various phenotypic markers were PE labeled as indicated. Select histograms from either HIV+ or HIV– patients were drawn based on gating the lineage-negative, HLA-DR+, DC-SIGN+ cells, and each phenotypic marker (thick line) was overlaid against the isotype-matched control antibody (thin line). Histogram plots below the thick line show the markers undetectable on the DC-SIGN+ cells. DC-SIGN+ cells didn't express langerin (data not shown). The two histograms on the left were drawn based on the lineage-positive gate to show detectable CCR5 (47% positive) and CXCR4 (60% positive) expression in the lineage-positive mucosal cell population. (C) CD14 and DC-SIGN colocalization. Rectal mucosal biopsy sections from a patient with unusually high numbers of CD14+ cells (red, left panel) are shown for illustrative purposes. Note that the majority of DC-SIGN+ cells (green, middle panel) are also CD14+ (yellow, right panel), consistent with the FACS data in panel B.

Primary gut mucosal DC-SIGN⁺ cells bind and transfer HIV-1. HIV-1 and simian immunodeficiency virus (SIV) have been shown to bind DC-SIGN. To characterize the virus binding and transfer ability of primary gut DCs, we obtained gut mucosal biopsies by flexible sigmoidoscopy. Up to 20 biopsies from a given patient were pooled and treated with collagenase to generate an MMC suspension. Due to the limited number of DC-SIGN⁺ cells that could be purified, we designed an ultrasensitive real-time RT-PCR assay to quantitatively measure the number of viral genomes bound to such a small population of cells. The calculated values were normalized to a standardized amount of ß-actin mRNA. Preliminary studies indicated that SIV Env-pseudotyped viruses gave the most robust binding signal and, thus, SIV316-pseudotyped viruses were first used to optimize this new binding assay.

First, we sorted total MMCs into HLA-DR⁺/DC-SIGN⁺ and HLA-DR⁺/DC-SIGN^– populations and bound virus in the presence or absence of various DC-SIGN inhibitors (Fig. 2A). In the three subjects examined, mannan and an anti-DC-SIGN antibody blocked virus binding by an average of 40 to 50%, indicating that part of the binding interaction was indeed specific to DC-SIGN (Fig. 2A). The level of virus capture by the DC-SIGN⁺ cells was in the range of 5 to 23 virions per 100 ß-actin mRNA copies. A low level of virus binding was also seen in the HLA-DR⁺/DC-SIGN^– population, but this binding was not inhibitable by mannan or the anti-DC-SIGN antibody.

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FIG. 2. Virus binding to DC-SIGN+ mucosal cells and transfer to T cells. (A) SIV316-pseudotyped pNL-GFP viruses were used in binding to sorted populations of HLA-DR+/DC-SIGN+ and HLA-DR+/DC-SIGN– gut cells from three different patients. HIV+ (, ) and HIV– (•) patient samples are indicated. The sorting gates were drawn as shown in Fig. 1A. The number of viral genomes bound per standard ß-actin mRNA copy number in the HLA-DR+/DC-SIGN+ population in the absence of any pretreatment (virus only) was normalized to 100%. (B) The total mucosal cell suspension was exposed to HIV-1 JR-CSF, and the DC-SIGN+ and DC-SIGN– populations were sorted from the HLA-DR+ gate. An aliquot of the unsorted virus-exposed MMCs was set aside. The amount of virus bound per ß-actin message under each condition was normalized to that bound to the HLA-DR+/DC-SIGN+ population (set at 100%). HIV+ () and HIV– (filled symbols) patient samples are indicated. (C) Total MMCs were exposed to HIV-1 JR-CSF with or without the DC-SIGN blocking antibody (clone 612 at 10 µg/ml), and the DC-SIGN+ and DC-SIGN– populations were sorted from the HLA-DR+ gate. The amount of virus bound per ß-actin message under each condition was normalized to that bound to the HLA-DR+/DC-SIGN+ population (set at 100%). The RT-PCR was run in quadruplicate and the error bars are shown, with asterisks denoting the DC-SIGN antibody, which significantly blocked virus binding. Results shown are for DC-SIGN+ cells (gray bars), DC-SIGN+ cells with blocking antibody (horizontal lined bars), DC-SIGN– cells (black bars), DC-SIGN– cells with blocking antibody (vertical and horizontal lined bars), MMCs (white bars), and MMCs with blocking antibody (vertical lined bars). (D) HIV-1 JR-CSF was titrated on peripheral blood MDDCs from 158 virions down to 1.58 virions/MDDC at the limit of detection. Five independent experiments are shown in circles, with the horizontal line indicating the average amount of HIV-1 virions detected per million ß-actin messages. (E) At decreasing viral inocula, virus was added to MDDCs pretreated as indicated and the amount of virus bound was expressed as a percentage relative to the level of virus bound under the untreated (virus-only) condition. Inocula were 158 virions/MDDC (gray bars) and 15.8 virions/MDDC (black bars). A representative experiment of two is shown. (F) MMCs were exposed to JR-CSF, and DC-SIGN+ and DC-SIGN– populations were sorted from the HLA-DR gate. Equal numbers of each sorted population were combined with day 2 phytohemagglutinin-stimulated CD4+ T cells at a 1:2 ratio. Supernatant was collected at days 1, 4, and 7, and a p24 enzyme-linked immunosorbent assay was performed to measure virus replication. Another transfer assay using a single-round SIV316-pseudotyped NL4-3-GFP virus also showed three- to fourfold greater transfer of virus in the DC-SIGN+ population over the DC-SIGN– population (data not shown).

To better model HIV-1 transmission, we exposed HIV-1 to the total MMC population and then sorted the HLA-DR⁺/DC-SIGN⁺ and HLA-DR⁺/DC-SIGN^– populations (Fig. 2B). In six different subjects, the HLA-DR⁺/DC-SIGN⁺ population bound about 15-fold more virus than the HLA-DR⁺/DC-SIGN^– population (Fig. 2B). More strikingly, the DC-SIGN⁺ population bound on average 40-fold more virus (per ß-actin signal) than the total MMC population (Fig. 2B). Thus, greater than 90% of virus bound to rectal MMCs was attributable to the DC-SIGN⁺ population. Calculations indicated that the DC-SIGN⁺ population captured 2.5 to 15 virions per 100 ß-actin messages, showing that similar levels of virus were bound when virus was exposed either to the total MMC population or just to the sorted DC-SIGN⁺ population alone (5 to 23 virions per 100 ß-actin messages) (see previous paragraph). Importantly, HIV-1 binding to the DC-SIGN⁺ population was more effectively blocked by DC-SIGN antibodies when the viral input was limiting. Thus, when the virus input was diluted up to 25-fold, virus binding appeared increasingly DC-SIGN specific, as indicated by increased blocking by an anti-DC-SIGN antibody from 50 to 60% to greater than 80% (P < 0.046 and P < 0.0004, respectively, compared to the unblocked control). Blocking by the same antibody wasn't consistently significant in the DC-SIGN^– or total mucosal cell population (P < 0.095 and P < 0.19, respectively) (Fig. 2C). The lowest amount of virus input used here is more consistent with the viral load found in seminal fluid (see Discussion).

We believe the experimental conditions used were within the linearity and sensitivity of our assay. Using the published value of 15,800 virions/pg of p24 (34), we titrated the number of virions added per MDDC from 158 virions/MDDC to 1.58 virions/MDDC and found that our assay resulted in a linear binding curve (Fig. 2D). Interestingly, as with the primary gut DC-SIGN⁺ cells, mannan and anti-DC-SIGN antibodies were better able to block virus binding at lower viral inocula (Fig. 2E). Figure 2E shows that while mannan and anti-DC-SIGN antibodies did not block virus binding at 158 virions/MDDC, they blocked virus binding by 40 to 60% when the virus inoculum was lowered 10-fold. Neither CD4 antibodies nor control mouse IgG blocked virus binding.

Next, we examine the ability of DC-SIGN⁺ cells in the gut to transfer virus to CD4⁺ T-cell blasts. To more closely model the cell populations that would be encountered by HIV during sexual mucosal transmission, virus was exposed to total MMCs and then the MMCs were sorted into CD3^–/HLA-DR⁺/DC-SIGN⁺ and CD3^–/HLA-DR⁺/DC-SIGN^– populations. Equal numbers of the sorted cells from each population were subsequently added to CD4⁺ T-cell blasts. Figure 2F shows that HLA-DR⁺/DC-SIGN⁺ cells clearly transferred more virus to the permissive T cells than the HLA-DR⁺/DC-SIGN^– cells. A fourfold difference in p24 production was already apparent by day 4.

DC-SIGN expression levels in the gut correlate with the mucosal IL-10/IL-12 ratio. Since our data indicated that DC-SIGN was at least a marker for gut cells with a DC phenotype that can bind and transfer virus, we next determined what effect established HIV-1 infection might have on DC-SIGN⁺ cells in the gut. We obtained rectal mucosal biopsy tissue sections from 26 HIV⁺ patients and 4 HIV^– healthy volunteers and detected DC-SIGN⁺ cells by immunofluorescence. The number of DC-SIGN⁺ cells in each section was quantified by computer-assisted morphometry. We found greater variability of DC-SIGN⁺ cells per standard area in HIV⁺ compared to HIV^– patients (HIV⁺ [29.4 to 154.6] versus HIV^– [22 to 63]) (Fig. 3). In addition, HIV⁺ patients had greater numbers of DC-SIGN⁺ cells per standard area than healthy HIV^– volunteers (73.1 ± 4.9 [HIV⁺, 42 sections from 26 individuals] versus 46.2 ± 5.6 [HIV^–, 8 sections from 4 individuals]) (mean ± standard error of the mean [SEM]; P < 0.0017) (Fig. 3). This difference appeared to be accounted for by a subset of HIV⁺ patients with high DC-SIGN⁺ counts, and it is possible that the significance of this difference may diminish when sections from greater numbers of HIV^– volunteers are counted.

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FIG. 3. An increase in DC-SIGN+ cells in the gut mucosa of HIV+ patients. Gut mucosal tissue sections were stained for DC-SIGN and detected by immunofluorescence. The DC-SIGN+ cells were enumerated by computer-assisted quantitative morphometry. Eight sections from 4 HIV– patients and 42 sections from 26 HIV+ patients were counted. The panels on the left show representative examples of patient samples with low, medium, and high numbers of DC-SIGN+ cells per standard area (defined as a 20x high-power field). The right panel is a graphical representation of the morphometric data obtained. At least five fields were counted per section; the data bar shown is the average count ± the SEM. The averages for all the HIV+ and HIV– sections (± SEM) were calculated (red bars) and compared using Student's t test (two-tailed unequal variance).

Since Th2 cytokines and/or a type 2 environment have been reported to upregulate DC-SIGN expression (36, 45), we sought to sample the immunological environment that might have given rise to the varied DC-SIGN counts that we observed. We restricted our analysis to the HIV⁺ patients, as the variance and the numbers in the HIV^– pool were not large enough for us to make any meaningful correlates. Thus, we measured the mRNA levels of the cytokines IL-10, IL-12 p40, IL-4, and TNF- by real-time RT-PCR in the same tissue used for immunofluorescent staining. Due to the variability of the absolute cytokine levels between each patient sample, it was difficult to assess a Th1 versus Th2 pattern between each patient. However, a good indicator of the overall Th2/Th1 balance is the ratio of IL-10 and IL-12 (24). By plotting the IL-10/IL-12 ratio versus DC-SIGN count, we found a strong positive correlation of the IL-10/IL-12 ratios and DC-SIGN cell counts in the gut (r = 0.58, P < 0.002; n = 26) (Fig. 4).

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FIG. 4. The mucosal IL-10/IL-12 ratio is positively correlated with increased numbers of DC-SIGN+ cells in mucosal tissue sections from HIV+ patients. Quantitative real-time RT-PCR was performed for IL-10 and IL-12 on RNA isolated from most of the HIV+ tissue sections, which were also quantified for DC-SIGN expression in Fig. 3. The log of the ratio of IL-10/IL-12 (x axis) was plotted against the DC-SIGN count (y axis) for each tissue section. A Pearson's correlation, r, the 95% confidence interval (bounded by the dotted lines), and P value of the correlation were determined by using the GraphPad Prism software at r = 0.58 and P = 0.0020.

To determine the effect of HIV infection on the phenotype of the DC-SIGN⁺ cells, we compared the phenotype of six HIV⁺ patients to that of six HIV^– patients. Irrespective of HIV status, we found that DC-SIGN expression was negatively correlated to the level of costimulatory molecules (CD80 and CD86) on the cognate DC-SIGN⁺ cells (CD86, r = –0.82, P < 0.002, n = 12; CD80, r = –0.81, P < 0.003, n = 11) (Fig. 5A and B, respectively). Figure 5C shows contrasting examples of patients with high and low numbers of DC-SIGN-expressing cells with their corresponding levels of CD80/CD86 expression. Thus, a shift towards a type 2 cytokine environment favored an increase in the number of DC-SIGN⁺ cells in the rectal mucosa, and an increased number of DC-SIGN⁺ cells was itself correlated with a decrease in the expression of the costimulatory molecules CD86 and CD80 on the DC-SIGN⁺ mucosal cells.

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FIG. 5. DC-SIGN+ cell frequency is negatively correlated with the mean fluorescent intensity (MFI) of the B7 family of costimulatory molecules. The percentage of DC-SIGN+ cells in the rectal MMC suspension was determined by flow cytometric gating on the lineage-negative, HLA-DR+, DC-SIGN+ cells in six HIV+ patients () and six HIV– patients (•) (y axis). The MFI of the B7 costimulatory molecules CD86 (A) and CD80 (B) on the lineage-negative, HLA-DR+, DC-SIGN+ cells was determined by subtracting the MFI of the isotype control staining from the MFI of either CD86 or CD80 (x axis). The Pearson's correlation, r, the 95% confidence interval (bounded by the dotted lines), and P value of the correlation were determined by using the GraphPad Prism software at r = –0.82 and P < 0.002 (A) and r = –0.81 and P < 0.003 (B). (C) Representative figure of how the data were collected for two points on the graph in panel A, representing high DC-SIGN frequency (5.8%) and low CD86 expression (MFI of 56) (top two panels) and low DC-SIGN frequency (0.8%) and high CD86 expression (MFI of 595) (bottom two panels).

IL-10 increases DC-SIGN, decreases CD86 expression, and functionally suppresses MDDCs. Based on the phenotypic differences we observed in vivo, we next sought to determine if IL-10 was a causal factor in both increasing DC-SIGN expression and decreasing costimulatory molecule expression in in vitro MDDC cultures. Rectal mucosal DCs potentially differentiate from their precursors once they have emigrated to the mucosa, and our preliminary data indicated that modulating the IL-10/IL-12 ratio in the MDDC cultures once differentiation had occurred (on day 5 to 7) had no effect on DC-SIGN expression intensity (data not shown). However, since the IL-10/IL-12 ratio could potentially influence DC precursors as they develop into DCs in the tissues, we decided to modulate the IL-10/IL-12 ratio while monocytes differentiated into DCs in the presence of IL-4 plus GM-CSF. Thus, IL-10 was added along with IL-4 plus GM-CSF from day 0 during MDDC differentiation. Figure 6A shows that MDDCs derived in the presence of IL-10 had greater DC-SIGN expression than those derived in the absence of additional IL-10 (compare immature DCs or mature DCs with immature IL-10-derived DCs and mature IL-10-derived DCs). This is unique to IL-10 in that addition of a similar amount of Th2 cytokines such as IL-4 and IL-13 did not induce the high levels of DC-SIGN expression seen on IL-10-derived MDDCs (data not shown). However, IL-10 could not replace the Th2 cytokines IL-4 or IL-13 for DC development, as monocytes cultured without IL-4 or IL-13 (i.e., GM-CSF plus IL-10) failed to differentiate into DCs (data not shown). In addition to the upregulation of DC-SIGN, MDDCs derived in the presence of IL-10 (with IL-4 plus GM-CSF) also resulted in DCs with decreased CD86 expression (Fig. 6A). Thus, IL-10 plays a causative role in the correlates we found on primary rectal mucosal DC-SIGN⁺ cells in vivo (Fig. 6A).

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FIG. 6. MDDCs in the presence of IL-10 upregulate DC-SIGN expression, downregulate CD86 expression, and fail to mature and migrate to appropriate stimuli. (A) Monocytes were cultured with IL-4 (100 ng/ml) plus GM-CSF (50 ng/ml) with or without IL-10 (100 ng/ml) for 7 days. Half the cells were stimulated with LPS (10 ng/ml) and TNF- (100 ng/ml) to induce maturation, and the other half was maintained in original cytokines in an immature state for 2 days. The expression levels of various markers were determined using CD86-TC, DC-SIGN-FITC, CD83-PE, and HLA-DR-APC by flow cytometric analysis, and the change in mean fluorescent intensity (MFI) was calculated by subtracting the MFI of the isotype control staining from the MFI of the cell surface marker indicated. The error bars are derived from triplicate cultures of one patient as a representative figure of five culture experiments. Results shown are for immature MDDCs (black bars), mature MDDCs (vertical and horizontal lined bars), immature IL-10-derived MDDCs (gray bars), mature IL-10-derived MDDCs (horizontal bars). (B) MDDCs were exposed to 250 ng/ml of MIP3-ß separated by a 0.5-µm polycarbonate membrane in a transwell system for 3 to 4 h. Cells that passed through were counted on a flow cytometer and expressed as the percent migration compared to medium only (no cytokine). The error bars represent the cumulative migration from four separate experiments. Mature MDDCs migrated to the MIP-ß despite undetectable levels of CCR7 expression by flow cytometric analysis.

As IL-10 is known to be an autocrine factor that influences DC development (9), we attempted to address the functional consequence of an elevated IL-10 environment. MDDCs derived with IL-10 were impaired in their ability to respond to maturation signals such as bacterial LPS and TNF- in that the mature DC marker CD83 and CD86 expression remained low relative to maturation induced on MDDCs derived without IL-10 (Fig. 6A). Effective maturation enables DCs to migrate via CCR7 to a MIP-3ß gradient in the secondary lymph node (12). Indeed, Fig. 6B shows that IL-10-treated DCs were significantly compromised in their ability to migrate towards a MIP-3ß gradient. Thus, the mechanism for DC-SIGN⁺ cell accumulation in the gut mucosa in the presence of higher levels of IL-10 may be a result of decreased DC migration away from the peripheral tissue to the secondary lymph nodes. In conclusion, an increased mucosal IL-10 environment correlates with a less immunostimulatory DC phenotype, which may contribute to the decrease in the overall immune function seen in HIV infection (25, 30).

Top Abstract Introduction Materials and Methods Results Discussion References

 
We found that DC-SIGN⁺ cells in the rectal mucosa comprise 1 to 5% of the cells in total mucosal cell suspensions (Fig. 1). Indeed, examining the immunofluorescent images revealed the abundance DC-SIGN⁺ cells in human rectal mucosal sections (Fig. 1C). DC-SIGN⁺ cells are also reportedly most abundant in this section of the gastrointestinal tract in rhesus macaques (23). DC-SIGN⁺ cells in the human rectal mucosa express markers associated with DCs (i.e., CD86, CD80, CD40, BDCA-3, and BDCA-4). Though typically associated with macrophages, CD14 expression is also detected on the rectal DC-SIGN⁺ population by both flow cytometry and immunofluorescence. This is also observed in the few rare DC-SIGN⁺ cells found in the blood (14). Bell et al. reported 0.6% of the colonic mucosal cell population are DCs, but they excluded CD14 in their definition of DCs and may have underestimated the total number of DCs (5). In our hands, a majority of DC-SIGN⁺ cells in the rectal mucosa also express CD14 and, thus, we may be characterizing a different DC type. More recently, te Velde et al. described two populations of DCs in the mucosa, one being CD83⁺ and one being DC-SIGN⁺, with differential expression of cytokines in patients of Crohn's disease (51). This is consistent with our present and previous studies (45), which found that DC-SIGN⁺ cells are generally CD83 negative. Here, we focused particularly on the DC-SIGN⁺ cells in the human rectal mucosa, due to their potential relevance for the capture and transmission of HIV.

Rectal mucosal DC-SIGN⁺ cells may be qualitatively different from peripheral blood DCs or MDDCs on the basis of CCR5 and CXCR4 expression, because the latter express easily detectable levels of CXCR4 and CCR5 with the same monoclonal antibody clones (27). We specifically note that these mucosal DC-SIGN⁺ cells appeared negative for the two major coreceptors, CCR5 and CXCR4, even though CXCR4 and CCR5 were readily detected on the lineage-positive cells from the same sample (Fig. 1B). Our results are discrepant from those of Jameson et al., who used triple-color confocal microscopy to show CCR5⁺, CD4⁺, and DC-SIGN⁺ staining from tissue sections of both humans and rhesus macaques (23). However, it was unclear what percentage of cells expressed both coreceptor and DC-SIGN; our isolation procedure and different sensitivities of the detection assays employed may also contribute to the discrepancies observed. Nevertheless, we speculate that the low or nonexistent expression of coreceptors on these DC-SIGN⁺ gut mucosal cells may lead to a more predominant role for DC-SIGN in the transfer of HIV from the periphery to T cells abundant in the secondary lymphoid organs, or to the abundant local CD4⁺, CCR5⁺ T cells that support viral replication in both acute and chronic phases of HIV disease (6, 29).

HIV-1 Env or virion binding and transmission studies have generally used in vitro-derived DCs from monocyte precursors or primary DCs isolated from the blood and skin. In this study, we characterized HIV-1 binding and transmission on the relevant primary rectal mucosal DC-SIGN⁺ cells, which would be encountered in sexual transmission. We isolated primary DC-SIGN⁺ cells from rectal mucosal biopsies by FACS using an antibody to the DC-SIGN repeat region (4), so as to not obstruct the virus binding site on the carbohydrate recognition domain of DC-SIGN. Due to the limiting numbers of cells obtained after cell sorting, we developed a sensitive and quantitative real-time RT-PCR assay to measure the number of genomic RNA copies bound per unit of mRNA for the housekeeping gene ß-actin.

Virus binding to sorted primary rectal mucosal DC-SIGN⁺ cells was partially blocked by excess mannan or an anti-DC-SIGN antibody (50%) (Fig. 2A), similar to what has been reported with MDDCs (3, 20, 52). However, Trumpfheller et al. observed more efficient blockade when using more than 2 logs less of virus (300 pg of p24) than what we have used in our initial experiments (52). To address this, we titrated the amount of virus on the total MMC population with or without a DC-SIGN blocking antibody. We found that at low viral inocula (e.g., 500 pg of p24), an anti-DC-SIGN antibody blocked virus binding to rectal mucosal DC-SIGN⁺ cells by almost 90% (Fig. 2C). This observation is of paramount importance, as this viral inoculum used is much closer to the viral load found in seminal fluid (from untreated HIV⁺ patients) (58) and almost 100-fold lower than the amount used in a previous study that showed no blocking of virion binding to MDDCs using mannan or anti-DC-SIGN antibodies (20). Thus, during sexual transmission, DC-SIGN could potentially be the critical player in the capture of HIV-1 and therefore is a potential target for therapeutic intervention to reduce viral transmission.

Strikingly, when virus was exposed to the total MMC population, about 40-fold more virus was bound to the DC-SIGN⁺ population compared to the total gut mononuclear cell population (Fig. 2B). Thus, greater than 90% of the bound virus was associated with the DC-SIGN⁺ cells, which constitute only 1 to 5% of the total MMC population. DC-SIGN serves at least as a marker of the cell type responsible for most of the virus binding and transfer activity present in MMCs.

We next asked what effect HIV-1 infection has on the DC-SIGN⁺ cells in the gut. Using quantitative morphometry on immunofluorescent-stained tissue sections, we counted the number of DC-SIGN⁺ cells per standard area. A subset of HIV-1⁺ patients had a two- to fourfold greater number of DC-SIGN⁺ cells infiltrating the lamina propria of the gut (Fig. 3). DC-SIGN⁺ cells are also increased in the colon of Crohn's disease patients (51) and SIV-infected macaques (8). We found that an increase in DC-SIGN expression in the gut mucosa correlated with a type 2 environment (increased IL-10/IL-12 ratio) and a decrease in the levels of the costimulatory molecules CD86 and CD80 (Fig. 5), regardless of HIV-1 infection status. IL-10 is known to be an immunosuppressive cytokine, and increased levels of IL-10 have also been correlated with other chronic infections, such as malaria, leprosy, tuberculosis, leishmaniasis, filariasis, and candidiasis (31). In the gut, IL-10 is crucial for the development of T_R1 regulatory T cells, which prevent colitis (19), probably via interaction with tolerogenic DCs generated in the presence of IL-10 (48). Although IL-10 treatment of in vitro MDDCs is known to give rise to DCs with a tolerogenic phenotype (9, 57) (Fig. 6), we have provided in vivo correlative data suggesting that IL-10 may also favor the development of tolerogenic DCs in the gut. Specifically, we make the novel observation that increased DC-SIGN expression in the gut (likely induced by increased IL-10 levels) is inversely correlated with CD80/CD86 expression and, thus, we implicate increased DC-SIGN expression as an additional marker for tolerogenic DCs. Our results are underscored by very recent results from microarray analysis experiments that IL-10-induced DCs indeed result in a DC-SIGN^high-expressing subset (56).

Mycobacteria take advantage of the immune-instructive capacity of DCs by signaling through DC-SIGN to secrete IL-10 and dampen the immune response (18). It is not know if HIV-1 could signal in the same manner; however, various HIV-1 proteins have been reported to induce IL-10 secretion in peripheral blood mononuclear cells (42, 50), and a recent report suggest that gp120 induced abnormal maturation of DCs that lack allostimulatory capacity (15). It is also intriguing to note that different polymorphisms in the promoter region of IL-10 have been linked to accelerated or decreased progression to AIDS (44). Thus, what virologic or immunologic factors influence the IL-10/IL-12 axis and DC-SIGN and how this affects the chronic viral reservoir in the gut are worthy of further investigation.

As a correlate to gut DCs, we used MDDCs to study the effects of the IL-10/IL-12 ratio on DCs and found that we could recapitulate our in vivo correlates: that IL-10 can increase DC-SIGN expression and decrease costimulatory molecule expression. Autocrine IL-10 produced by the MDDCs prevents spontaneous maturation (9); thus, preventing maturation prevents the maturational-induced decrease of DC-SIGN expression (39). Not only would the IL-10-derived DCs be blocked in spontaneous maturation, but also experimentally induced maturation by bacterial LPS and TNF- was also impaired by IL-10 (Fig. 6A). Thus, the addition of IL-10 may direct MDDC development to a hyper-immature state with greater DC-SIGN expression levels. Immature DCs express the chemokine receptor CCR6 for migrating to MIP-3 expressed in peripheral tissues and, upon maturation of DCs, CCR7 expression increases for trafficking to MIP-3ß expression in the secondary lymph nodes (16, 38). Just as IL-10 treatment also influences multiple transcriptional programs, such as those involving chemotaxis (11, 33, 35), we also found that derivation of DCs in the presence of IL-10 impaired chemotaxis to MIP-3ß in vitro. Indeed, murine DCs derived in vitro with IL-10 downregulated CCR7 and had decreased in vivo homing ability (49). Thus, the increased IL-10 levels in the gut microenvironment may maintain the resident DCs in an immature CCR6⁺/CCR7^–-expressing state, thereby limiting emigration from the peripheral tissues, leading to an accumulation in the gut tissue.

To our knowledge, this is the first demonstration that relevant rectal mucosal DC-SIGN⁺ cells can bind and transfer HIV to permissive T cells. We also provide in vivo and ex vivo data that suggest a close relationship between DC-SIGN and costimulatory molecule (CD80/CD86) expression on DCs—this relationship is functionally modulated by IL-10 levels. Our data suggest the nexus of IL-10 modulation with DC-SIGN and costimulatory molecule expression on DCs could be a vital part of the viral immune evasion strategy and is worth further experimental investigation. In summary, we have defined a cell population in the human rectal mucosa that plays a critical role in the virus-host interaction, and we have characterized the in situ parameters that might modulate the function of these cells. Our data also provide fresh insight into the dynamics of mucosal DC populations.

 
K.B.G. is supported by a Ruth Kirschstein Postdoctoral fellowship. B.L. is a Charles E. Culpepper Medical Scholar supported by the Rockefeller Brothers Fund and a recipient of the Burroughs Wellcome Fund Career Development Award, and he is supported by NIH grants RO1-AI52021 and R21-AI055305. We also acknowledge support of the UCLA AIDS Institute and the flow cytometry, virology, and mucosal immunology cores (UCLA CFAR grant NIH AI-28697) and the James B. Pendleton Charitable Trust. P.A.A. is supported by NIH grants RO1-AI50467, K24 AI01610, and AI28697 as well as the Macy's Foundation and the Oppenheimer Brothers' Foundation.

We thank Jerry Zack and Jon Braun for constructive criticisms of the manuscript. We also thank Marie Fuerst for coordination of the patient scheduling and our patients and subjects for volunteering rectal mucosal biopsies.

 
* Corresponding author. Mailing address for P. A. Anton: Dept. of Medicine, Division of Digestive Diseases, David Geffen School of Medicine at UCLA, 2734 MRL, 675 Charles E. Young Dr. South, Los Angeles, CA 90095-7019. Phone: (310) 206-5797. Fax: (310) 206-8824. E-mail: Panton{at}mednet.ucla.edu.

* Corresponding author. Mailing address for B. Lee: Dept. of Microbiology, Immunology & Molecular Genetics, David Geffen School of Medicine at UCLA, 3825 Molecular Sciences Building, 609 Charles E. Young Drive East, Los Angeles, CA 90095-1489. Phone: (310) 794-2132. Fax: (310) 267-2580. E-mail: bLeebhL{at}ucla.edu.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Journal of Virology, May 2005, p. 5762-5773, Vol. 79, No. 9
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.9.5762-5773.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

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