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Originally published In Press as doi:10.1074/jbc.M503401200 on July 28, 2005

J. Biol. Chem., Vol. 280, Issue 39, 33123-33131, September 30, 2005
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PU.1 Regulates the Tissue-specific Expression of Dendritic Cell-specific Intercellular Adhesion Molecule (ICAM)-3-grabbing Nonintegrin*

Ángeles Domínguez-Soto1, Amaya Puig-Kröger2, Miguel A. Vega, and Angel L. Corbí3

From the Centro de Investigaciones Biológicas, CSIC, Madrid 28040, Spain

Received for publication, March 29, 2005 , and in revised form, June 13, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Dendritic cell-specific intercellular adhesion molecule-3-grabbing nonintegrin (DC-SIGN) is a cell surface C-type lectin expressed on myeloid dendritic cells and certain tissue macrophages, which mediates antigen capture for processing and presentation and participates in intercellular interactions with naive T lymphocytes or endothelial cells. In their strategy to evade immunosurveillance, numerous pathogenic microorganisms, including human immunodeficiency virus and Mycobacterium, bind to DC-SIGN in order to gain access to dendritic cells. We present evidence that PU.1 dictates the basal and cell-specific activity of DC-SIGN gene-regulatory region through in vivo occupancy of two functional Ets elements, whose integrity is required for PU.1 responsiveness and for the cooperative actions of PU.1 and other transcription factors (Myb, RUNX) on the DC-SIGN gene proximal regulatory region. In addition, protein analysis and gene profiling experiments indicate that DC-SIGN and PU.1 are coordinately expressed upon classical and alternative macrophage activation and during dendritic cell maturation. Moreover, small interfering RNA-mediated reduction of PU.1 expression results in diminished DC-SIGN cellular levels. Altogether, these results indicate that PU.1 is involved in the myeloid-specific expression of DC-SIGN in myeloid cells, a contribution that can be framed within the role that PU.1 has on the acquisition of the antigen uptake molecular repertoire by dendritic cells and macrophages.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Dendritic cell-specific intercellular adhesion molecule-3-grabbing nonintegrin (DC-SIGN4; CD209) (1) is a type II membrane protein and C-type lectin whose carbohydrate-binding domain is followed by a "neck" region with seven 23-amino acid repeats, a transmembrane region, and a short cytoplasmic tail with recycling and internalization motifs (2, 3). DC-SIGN is preferentially expressed on myeloid dendritic cells (3, 4) and is usually considered as a dendritic cell (DC)-specific phenotypic marker. However, although DC-SIGN expression in vivo is mainly restricted to interstitial DC (5), it has also been detected on synovial, placenta, and alveolar macrophages and on a small subset of CD14+ peripheral blood DC (6–9). DC-SIGN expression is induced de novo by IL-4 during the in vitro generation of monocyte-derived dendritic cells (MDDC) (4), upon alternative activation of macrophages (7, 10) and during differentiation of THP-1 cells in the presence of protein kinase C activators (10).

Functionally, DC-SIGN mediates antigen capture for processing and presentation in the context of major histocompatibility complex class II molecules (11, 12), participates in the establishment of the initial contact between dendritic cells and naive T lymphocytes through its recognition of intercellular adhesion molecule-3 (3), and also mediates dendritic cell trafficking through interactions with endothelial intercellular adhesion molecule-2 (13). On the other hand, numerous reports have provided evidence that DC-SIGN function is subverted by numerous pathogens, and a large array of pathogenic microorganisms, including virus (HIV, cytomegalovirus, Ebola, hepatitis C, Dengue), bacteria (Mycobacterium tuberculosis, Helicobacter pylori), fungi (Candida, Aspergillus), and parasites (Leishmania, Schistosoma mansoni) (reviewed in Ref. 1), bind to DC-SIGN in a mannose- or fucose-dependent manner (14–17) as a strategy to escape from immunosurveillance. Subsequent to binding to DC-SIGN, most of these pathogens initiate their replicative or infective cycle from the cytoplasm of DC-SIGN-expressing cells. In the case of HIV, DC-SIGN mediates virus binding and uptake by dendritic cells, where DC-SIGN-associated HIV remains infectious for prolonged periods of time and efficiently transfers HIV to CD4(+) T cells in lymph nodes, where viral replication occurs (18). The critical function of DC-SIGN in HIV entry and spread is illustrated by the existence of DC-SIGN promoter polymorphisms associated with increased risk of parenteral infection by HIV-1 (19). Therefore, the identification of the regulatory sequences driving DC-SIGN basal and regulated expression might provide valuable information of potential diagnostic and therapeutic relevance.

PU.1 is the most divergent member of the Ets family of transcription factors whose expression is restricted to myeloid and B-lymphoid cells (20, 21). PU.1 contacts DNA with a winged helix-turn-helix-type domain, binds to a cis element (core sequence 5'-GAGGAA-3') that is sometimes clustered with binding sites for other transcription factors, and regulates the expression of genes involved in cytokine binding (macrophage colony-stimulating factor receptor and granulocyte colony-stimulating factor receptor), cell adhesion (CD11b, CD11c, CD18, and scavenger receptors), and components of the phagocyte NADPH-dependent oxidase system (20). PU.1 is required for terminal myeloid differentiation and gene expression, and myeloid and B lymphoid lineage cells either completely fail to develop (22) or are delayed and highly aberrant (23) in PU.1 –/– mice. Recent studies have demonstrated that different cellular concentrations of PU.1 may direct distinct cell fates, with the highest concentrations of PU.1 required for macrophage development and lower concentrations for granulocytic and B-cell fate adoption (24, 25). PU.1 is also involved in the development of myeloid-derived populations of murine dendritic cells (26, 27) and promotes dendritic cell development from CD34+ human cord blood cells and generation of Langerhans cells, an effect that is blocked by CCAAT/Enhancer-binding protein {alpha} (28). In the present report, we describe the coordinated expression of PU.1 and DC-SIGN during macrophage activation and dendritic cell maturation and the critical contribution of PU.1 to the basal and tissue-specific activity of the DC-SIGN gene proximal regulatory region through occupancy of two closely located binding sites.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture—Human peripheral blood mononuclear cells were isolated from buffy coats from normal donors over a Lymphoprep (Nycomed Pharma, Oslo, Norway) gradient according to standard procedures. Monocytes were purified from peripheral blood mononuclear cells by a 1-h adherence step at 37 °C in complete medium or by magnetic cell sorting using CD14 microbeads (Miltenyi Biotech, Bergisch Gladbach, Germany). To generate MDDC, adherent or CD14+ cells (>95% monocytes) were cultured at 0.5–1 x 106 cells/ml in RPMI supplemented with 10% fetal calf serum, 25 mM HEPES, and 2 mM glutamine (complete medium), at 37 °C in a humidified atmosphere with 5% CO2, and containing 1000 units/ml granulocyte-macrophage colony-stimulating factor (Leucomax, Schering-Plough, Kenilworth, NJ) and 1000 units/ml IL-4 (PreProtech, Rocky Hill, NJ) for 5–7 days, with cytokine addition every second day. For MDDC maturation, immature MDDC were treated with lipopolysaccharide from Escherichia coli 055:B5 (10 ng/ml) for 24–48 h. To generate monocyte-derived macrophages (MDM), adherent or CD14+ cells were cultured at 0.5–1 x 106 cells/ml in complete medium containing 1000 units/ml granulocyte-macrophage colony-stimulating factor (or macrophage colony-stimulating factor at 10 ng/ml) for 5 days, with cytokine addition every second day. MDM were then treated with IL-4 (1000 units/ml) or interferon-{gamma} (500 units/ml) for 48 h to generate alternatively activated macrophages (AAM{Phi}) or classically activated macrophages (CAM{Phi}), respectively.

The K562 (chronic myelogenous leukemia) and THP-1 (monocytic leukemia) cell lines were cultured in RPMI supplemented with 10% fetal calf serum, 25 mM HEPES, and 2 mM glutamine (complete medium) at 37 °C in a humidified atmosphere with 5% CO2. NIH-3T3 cells were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum. Induction of differentiation of THP-1 cells was accomplished in the presence of phorbol myristate acetate at 10 ng/ml. When indicated, IL-4 was added at 1000 units/ml. In differentiation experiments, cells were seeded at 5 x 105 cells/ml in tissue culture dishes with no change of the culture medium after the addition of the differentiation inducer, and differentiation was allowed to proceed for 48 h.

Transfections, Plasmids, and Site-directed Mutagenesis—Transfection in COS-7, NIH-3T3, Jurkat, and K562 cells was performed with Superfect (Qiagen, Hilden, Germany) according to the manufacturer's instructions. Transfections were carried in 24-well plates out using 1 µg of eukaryotic expression plasmid DNA on 4 x 104 (COS-7, NIH-3T3) or 1 µg of firefly luciferase-based reporter plasmid and 100–200 ng of eukaryotic expression plasmid DNA on 8–15 x 105 (K562) cells. THP-1 cells were transfected using DEAE-dextran following standard procedures. In reporter gene experiments, the amount of DNA in each transfection was normalized by using the corresponding insertless expression vector (CMV-0) as carrier. Each transfection experiment was performed at least three times with different DNA preparations. Transfection efficiencies were normalized by cotransfection with the pCMV-{beta}gal plasmid, and {beta}-galactosidase levels were determined using the Galacto-Light kit (Tropix, Bedford, MA). The DC-SIGN-based reporter gene constructs pCD209-1600 and pCD209–468 reporter plasmids were generated by PCR amplification of the –1656/–19 and –468/–19 fragments of the DC-SIGN promoter with oligonucleotides 5'-CCAAGCTTTGTCAACTCACTTTCAGTTATACGTC-3', 5'-CCAAGCTTCAATTATGATTCTGCCCCAACTCC-3' and 5'-GGGGTACCAGTGTCCAGAACTCCTGGGG-3', and the resulting fragments were cloned into HindIII and KpnI-digested pXP2 plasmid, which contains the firefly luciferase cDNA (29). The pCD209-1200 reporter plasmid was generated by digestion of pCD209–1600 plasmid with BamHI and subsequent religation. Positions within the DC-SIGN regulatory regions were numbered from the ATG initiation start codon (+1). The PU.1 expression plasmid pCDNA3.1-PU.1 was obtained by cloning of the human PU.1 cDNA into EcoRI-digested pCDNA3.1(–) and selection of plasmids with properly oriented inserts.

THP-1 cells (2 x 106 cells) were transfected by nucleofection with 3 µg of siRNA for PU.1 (sc-36330 PU.1 siRNA gene silencer; Santa Cruz Biotechnologies, Inc., Santa Cruz, CA) or a control siRNA using the Cell Line Nucleofector Kit V (Amaxa). After nucleofection, cells were maintained in culture for 24 h, and one-fifth of the cells were lysed and subjected to Western blot for detection of PU.1, DC-SIGN, or {beta}-actin as control. Total RNA was isolated from the rest of nucleofected cells and subjected to reverse transcription-PCR for detection of DC-SIGN and glyceraldehyde-3-phosphate dehydrogenase RNA, as previously described (10). 2 µg of RNA from cells transfected with either PU.1-specific or control siRNA was reverse transcribed, and 5 µl of the resulting cDNA was subjected to PCR with oligonucleotides CD209sense (5'-GGGAATTCAGAGTGGGGTGACATGAGTGAC-3') and CD209antisense (5'-CCCCAAGCTTGTGAAGTTCTGCTACGCAGGAG-3') to amplify the whole coding region of DC-SIGN. Control PCRs were performed using oligonucleotides 5'-GGCTGAGAACGGGAAGCTTGTCA-3' and 5'-CGGCCATCACGCCACAGTTTC-3', which together amplify a 417-bp fragment from the glyceraldehyde-3-phosphate dehydrogenase mRNA. PCR-generated fragments were resolved in 1.5% agarose gels.

Site-directed mutagenesis was performed on the DC-SIGN promoter construct pCD209–468 using the QuikChange system (Stratagene, La Jolla, CA). For mutation of the PU-1–111 and PU.1–77 elements, the oligonucleotides PU.1–111mutS (5'-GGATGACAGATCCCTACCCGTCGACCTGTTTCTCTTTCTGTGGG-3'), PU.1–111mutAS (5'-CCCACAGAAAGAGAAACAGGTCGACGGGTAGGGATCTGTCATCC-3'), PU.1–77mutS (5'-CTGTGGGAGACTAGATTGTCGACGTAAAGATCACAGGGTGGG-3'), and PU.1–77mutAS (5'-CCCACCCTGTGATCTTTACGTCGACAATCTAGTCTCCCACAG-3') were used, and the resulting plasmids were termed pCD-209–468–111MUT and pCD209–468–77MUT. All DNA constructs and mutations were confirmed by DNA sequencing. Generation of the pCD209–468–111/–77MUT plasmid, where the Ets-binding sequences at –111 and –77 are mutated, was accomplished by site-directed mutagenesis on the pCD209–468–77MUT plasmid using the oligonucleotides PU.1–111mutS and PU.1–111mutAS. Mutation of the T/C-rich region downstream from the PU.1–111 element was accomplished using the oligonucleotides CD209–105mutS (5'-CCTACCCAACTTCCTGTTTGTCGACCTGTGGGAGACTAGATTTAG-3') and CD209–105mutAS (5'-CTAAATCTAGTCTCCCACAGGTCGACAAACAGGAAGTTGGGTAGG-3'), and the resulting plasmid was termed pCD209–468–105MUT.

Electrophoretic Mobility Shift Assays (EMSA)—Nuclear extracts were prepared according to Schreiber et al. (30), and EMSA was performed as described (31, 32). Unlabeled competitor oligonucleotides were added to the nuclear extracts at a 100-fold molar excess and incubated at 4 °C for 15 min before the addition of the radioactive probe. In inhibition/supershift experiments, 1 µl of anti-human PU.1 polyclonal antiserum (reference or origin) was incubated with the nuclear extract before the addition of the probe. Oligonucleotide probes used for EMSA were DC-SIGN-1 (5'-GTGGGAGACTAGATTTAGGAAGTAAAGA-3', spanning the DC-SIGN promoter between –94 and –67) and DC-SIGN-2 (5'-TACCCAACTTCCTGTTTCTCTTTCTGTGGGAGACT-3', based on the DC-SIGN promoter sequence –119/–85). Double-stranded oligonucleotides used as competitors included DC-SIGN-1, DC-SIGN-2, and DC-SIGN2mut (5'-GGATGACAGATCCCTACCCGTCGACCTGTTTCTCTTTCTGTGGG-3'), which includes the DC-SIGN promoter sequence between –119 and –85 mutated at the PU.1–111 binding site.

Western Blot—Cell lysates were obtained in 50 mM HEPES, pH 7.5, 250 mM NaCl, 1 mM EDTA, 0.5% Triton X-100, 0.5 mM dithiothreitol, 10 mM NaF, 1 mM Na3VO4, 2 mM Pefabloc, and 2 µg/ml aprotinin, antipain, leupeptin, and pepstatin. Nuclear and cytoplasmic extracts were prepared according to Schreiber et al. (30). 10 µg of cell lysates was subjected to SDS-PAGE under reducing conditions and transferred onto an Immobilon polyvinylidene difluoride membrane (Millipore Corp., Bedford, MA). After blocking of the unoccupied sites with 5% nonfat dry milk in 50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.1% Tween 20, protein detection was performed using the Supersignal West Pico Chemiluminescent system (Pierce). For reprobing, membranes were incubated in stripping buffer (62.5 mM Tris-HCl, pH 6.7, 100 mM {beta}-mercaptoethanol, 2% SDS) for 30 min at 50 °C with occasional agitation. Detection of PU.1 was carried out using specific affinity-purified rabbit polyclonal antibody (sc-352; Santa Cruz Biotechnology, Inc., Santa Cruz, CA). DC-SIGN was detected using a polyclonal antiserum raised against the 23-residue repeats within the neck region of the molecule (10).

Flow Cytometry and Antibodies—Phenotypic analysis was carried out by indirect immunofluorescence, using unlabeled primary monoclonal antibodies followed by incubation with fluorescein isothiocyanate-labeled Fab goat anti-mouse IgG. Monoclonal antibodies included HB1/5 (anti-CD83, Immunotech, Marseille, France), MR1 (anti-DC-SIGN, CD209), and Bear-2 (anti-CD14). All incubations were done in the presence of 50 µg/ml human IgG to prevent binding through the Fc portion of the antibodies. The supernatant from the myeloma P3X63Ag8 (X63) was always included as negative control. Flow cytometry analysis was performed with an EPICS-CS (Coulter Científica, Madrid, Spain) using log amplifiers.

Chromatin Immunoprecipitation—Chromatin immunoprecipitation assays were performed using the chromatin immunoprecipitation assay kit (Upstate%20Biotechnology">Upstate Biotechnology, Inc., Charlottesville, VA) following the manufacturer's instructions. MDDC cells were cross-linked with 1% formaldehyde for 5 min at 37 °C. After washing with ice-cold PBS, cells were lysed in 200 µl of a solution containing 1% SDS, 10 mM EDTA, 50 mM Tris-HCl, pH 8.1, and including 1 µg/ml aprotinin, leupeptin, and pepstatin and 1 mM phenylmethylsulfonyl fluoride. Chromatin samples were sonicated with four sets of 10-s pulses at 30% maximum power in a Sonifier S-150D to reduce DNA length to ~200–600 bp. Sonicated lysates were then diluted to 2 ml with 0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 167 mM NaCl, 16.7 mM Tris-HCl, pH 8.1, and 20 µl of this solution were removed for later PCR analysis (input). After preclearing with salmon sperm DNA/Protein A-agarose for 1 h at 4 °C, antibodies (2 µg) were added, and the sonicated lysates were incubated overnight at 4 °C in a rocking platform. Immune complexes were collected with 60 µl of salmon sperm DNA/Protein A-agarose (1 h at 4 °C), and agarose beads were washed with solutions of increasing ionic strength. After a final wash in 1 mM EDTA, 10 mM Tris-HCl, pH 8.0, bound immune complexes were eluted in a freshly prepared solution of 1% SDS, 0.1 M NaHCO3, and cross-links were reversed in the samples (and the input from the sonicated lysates) by heating at 65 °C for 4 h. Samples were then treated with Proteinase K, and DNA was phenol/chloroform-extracted and precipitated. DNA was resuspended (20 µl), and 2 µl was used for detection of the DC-SIGN promoter by PCR using the oligonucleotides 5'-TGCCTACCCTTGCCCTAGTGGA-3' and 5'-TGGAGTCACTCATGTCACCCCAC-3', which together amplify 353 bp spanning from –340 to +13. DNA from the input was resuspended in 20 µl, and 2 µl was used for PCR. Immunoprecipitating antibodies included affinity-purified rabbit polyclonal antibody against human PU.1 (sc-352; Santa Cruz Biotechnology) and purified polyclonal rabbit IgG as a control. As a positive control, amplification of the CD68 promoter was achieved with oligoncleotides 5'-CAACTGCCCTAGGACTCCGTTTG-3' and 5'-TCAGCCTCCTTCCTCCTTACCT-3', which together amplify a 318-bp region (33).

Gene Expression Profiling in Dendritic Cells—RNAs from immature and lipopolysaccharide-matured MDDC from two independent donors were labeled, processed, and independently hybridized on Codelink human whole genome DNA chip of the Codelink microarray platform (Amersham Biosciences), containing 55,000 human gene targets. For one of the donors, three replicates were independently hybridized. Scanned images were processed using the Codelink Expression Analysis software. Raw data were normalized by the quantile method. Data corresponding to the experimental groups (immature MDDC versus mature MDDC) was analyzed by Student's t test. The raw p values obtained were adjusted using the control of the false discovery rate-based procedure developed by Benjamini and Yekutieli (34) and implemented in the multitest package within the Bioconductor set of routines (available on the World Wide Web at www.bioconductor.org). Those genes with adjusted p values lower than 0.01 and greater than 2-fold differences in both donors were considered as differentially expressed.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Tissue-specific Activity of the DC-SIGN Promoter—Besides MDDC and {Delta}{Delta}M{Phi} (6–10), DC-SIGN expression can be seen on THP-1 myeloid cells but not on lymphoid or erythroleukemic cell lines like Jurkat or K562 cells (10, 35) (Fig. 1, A and B). To analyze the DNA elements and transcription factors that direct the expression of DC-SIGN, we amplified by PCR a genomic region of the DC-SIGN gene (–1656/–19) where, in agreement with a previous report (36), preliminary 5'-rapid amplification of cDNA ends experiments mapped the DC-SIGN transcriptional start sites (data not shown). Sequencing the amplified region revealed its complete identity to the genomic region upstream of the structural region of the DC-SIGN gene that had been determined previously in the contig sequence AC008812 [GenBank] .7.1.143619 within the Ensembl data base. Based on the sequence and the presence of numerous repetitive sequences in the distal region, two additional deletion constructs were generated, which span the regions –1275/–19 and –468/–19 to dissect the functional activity of the DC-SIGN promoter (Fig. 1C). To determine whether the DC-SIGN regulatory region exhibited differential activity in DC-SIGN+ and DC-SIGN– cells, its activity was assayed in myeloid THP-1, erythroleukemic K562, and lymphoid T Jurkat cell lines. The three DC-SIGN promoter-based constructs exhibited considerably higher activity in THP-1 than in Jurkat or K562 cells, and similar results were obtained when using NIH-3T3 cells (Fig. 1D and not shown). On average, the activity of the pCD209–1600Luc, pCD209–1200Luc, and pCD209–468Luc constructs was 75-, 140-, and 160-fold higher in the DC-SIGN+ THP-1 cell line than in nonexpressing cells (Fig. 1D). Therefore, it can be concluded that the DC-SIGN proximal promoter displays cell-specific activity and exhibits a higher activity in myeloid cells with constitutive expression of DC-SIGN and that most of the tissue-specific activity is retained in the proximal region of the promoter –468/–19.



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  		FIGURE 1.
Restricted expression of DC-SIGN and cell-specific activity of the DC-SIGN promoter. A, cell surface expression of DC-SIGN on THP-1, Jurkat, and K562 cell lines. Flow cytometry was performed using the MR-1 anti-DC-SIGN monoclonal antibody (4). B, determination of DC-SIGN expression in THP-1, Jurkat, and K562 cell lines by Western blot, using a polyclonal antiserum against the neck region of the molecule. C, schematic representation of the proximal regulatory region of the DC-SIGN gene and reporter plasmids used for its functional dissection. D, the DC-SIGN promoter-based constructs pCD209–1600, pCD209–1200 and pCD209–468 were transfected in THP-1 (DC-SIGN+), Jurkat (DC-SIGN–), and K562 (DC-SIGN–). After 48 h, cells were lysed, and luciferase activity was determined. In all cases, the pCMV-{beta}gal plasmid was cotransfected with the luciferase reporter constructs, and transfection efficiencies were normalized according to the {beta}-galactosidase levels. Data represent mean ± S.D. of four independent experiments using two different DNA preparations.

 
PU.1 Binds the DC-SIGN Promoter in Vitro and in Vivo—Initial gel shift assays on oligonucleotides spanning the –468/–19 and software-assisted prediction of transcription factor binding sites revealed the presence of the sequences 5'-CCCAACTTCCTGTTTC-3' and 5'-AGATTTGGAAGTAAAG-3' at –111 and –77, respectively, both containing the core recognition motif of the Ets family of transcription factors (Fig. 1C). The pattern of EMSA retarded complexes on oligonucleotides based on these sequences (oligonucleotide probes DC-SIGN-1 and DC-SIGN-2) revealed that they were specifically recognized by nuclear extracts from THP-1 and monocyte-derived dendritic cells but not by K562 cells (Fig. 2A). Moreover, the pattern of retarded complexes on both probes was similar, if not identical, suggesting that they were recognized by a common factor. The pattern of retarded complexes, their cell distribution, and the presence of the GGAA sequence led us to hypothesize that both elements were recognized by the PU.1 transcription factor. As shown in Fig. 2B, the integrity of the GGAA sequence within both oligonucleotides was absolutely required for complex formation. The demonstration that PU.1 binds both elements at the DC-SIGN promoter was obtained through the use of nuclear extracts from PU.1-transfected COS-7 cells and an anti-PU.1 polyclonal antiserum; the mobility of the major retarded complex was identical in PU.1-transfected cells, THP-1 and monocyte-derived dendritic cells, and, more importantly, the complexes were inhibited and supershifted in the presence of the anti-PU.1 polyclonal antiserum (Fig. 2B). The mutation introduced at the distal PU.1 element also affected the formation of a slow mobility complex whose formation was dependent on Sp1 (Fig. 2B). However, the ability of Sp1 to transactivate the DC-SIGN promoter was not affected by such a mutation (see below). Altogether, these results demonstrate that the PU.1 transcription factor interacts with two sequences within the proximal regulatory region of the DC-SIGN gene, one of which is located adjacent to an Sp1-binding site.

To confirm that PU.1 interacts with the proximal region of the DC-SIGN promoter, we performed chromatin immunoprecipitation with an anti-PU.1 antibody on human monocyte-derived dendritic cells, which exhibit a very high level of expression of DC-SIGN. The proximal DC-SIGN promoter region could be readily amplified from anti-PU.1 immunoprecipitated chromatin, whereas no amplification was obtained in the absence of antibody or in the presence of a control antibody (Fig. 3A). The CD68 promoter, whose occupancy by PU.1 has been previously demonstrated (33), was also amplified from the DNA immunoprecipitated by the anti-PU.1 antiserum (Fig. 3B). Therefore, PU.1 recognizes the proximal promoter of DC-SIGN in vitro, and its binding can also be detected in vivo by means of chromatin immunoprecipitation.

PU.1 Contributes to the Basal and Tissue-specific Activity of the DC-SIGN Proximal Regulatory Region—The relevance of PU.1 binding to the DC-SIGN promoter was analyzed in two alternative manners. First, the effect of independent mutations at each PU.1-binding site was evaluated in DC-SIGN+ and DC-SIGN– cells in the context of the –468/–19 region of the promoter. Mutation of the distal PU.1-binding element led to a dramatic reduction (only 5% of the activity of the wild type construct) in the DC-SIGN promoter activity, whereas disruption of the proximal PU.1 element reduced promoter activity by half (Fig. 4A). Moreover, the simultaneous mutation of the PU.1-binding elements at –111 and –77 resulted in a dramatic and very significant (p < 10–5) reduction of the DC-SIGN promoter activity (Fig. 4B), implying that both sites contribute to the basal activity of the DC-SIGN promoter. The reduction in activity caused by mutation of each PU.1-binding site was higher in DC-SIGN+ (THP-1) than in DC-SIGN– cells (Fig. 4A), indicating that both sites contribute to the tissue-specific activity of the promoter. As a control, mutation of the sequences at –105, predicted to be recognized by members of the IRF family, had a weaker effect in DC-SIGN+ cells.



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  		FIGURE 2.
Nuclear factors interacting with the DC-SIGN-1 and DC-SIGN-2 oligonucleotide probes and identification of specific nucleotides involved. A, EMSA was performed on the DC-SIGN-1 (left panel) and DC-SIGN-2 (right panel) oligonucleotides using nuclear extracts from K562 cells, immature MDDC, and THP-1 cells untreated or differentiated with phorbol 12-myristate 13-acetate + IL-4 (p + 4). The position of the major retarded species is indicated. Where indicated, unlabeled competitor oligonucleotides were added at a 100-fold molar excess. B, EMSA was performed on the DC-SIGN-2 oligonucleotide using nuclear extracts from THP-1 cells, immature MDDC or COS-7 cells transfected with an empty pCDNA3.1(–) expression vector or with the human PU.1 expression vector pCDNA3.1-PU.1, and in the absence or presence of the indicated competitor oligonucleotides at a 100-fold molar excess or an anti-PU.1 polyclonal antiserum. The positions of PU.1-containing complexes and the anti-PU.1-supershifted complex are indicated.

 



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  		FIGURE 3.
In vivo occupancy of the DC-SIGN proximal promoter by PU.1. Shown are chromatin immunoprecipitations on immature monocyte-derived dendritic cells using an affinity-purified polyclonal antiserum specific for PU.1, a nonspecific affinity-purified antiserum (control Ab), or no antibody. Immunoprecipitated chromatin was analyzed by PCR using a pair of DC-SIGN promoter-specific primers that amplify a 353-bp fragment flanking the PU.1-binding sites at –111 and –77 (A) or primers specific for detection of the CD68 promoter (B). Input lanes represent the PCR analysis performed on DNA from a 1:20 dilution of the starting sonicated lysate. Each experiment was performed twice with similar results, and one of the experiments is shown.

 
Second, the ability of PU.1 to transactivate the DC-SIGN promoter was determined in cells devoid of PU.1 expression (NIH-3T3). PU.1 was capable of transactivating, weakly but significantly, the DC-SIGN promoter when the reporter construct was co-transfected with a PU.1 expression vector (Fig. 4C), and the transactivating effect was reduced by either independent or simultaneous mutation of both PU.1-binding sites (Fig. 4C). Altogether, this set of results demonstrates that occupancy of the PU-1 binding sites contributes to both the basal and tissue-specific activity of the DC-SIGN promoter.

PU.1 Transactivates the DC-SIGN Proximal Regulatory Region in Collaboration with Other Myeloid Transcription Factors—To determine whether other myeloid factors play a role in the activity of the DC-SIGN promoter, we evaluated the influence of Myb and RUNX factors in transactivation experiments, since both are expressed in myeloid cells and have been previously reported to collaborate with PU.1 (37, 38). Preliminary experiments in K562 cells revealed that the DC-SIGN promoter was transactivated by RUNX1 (8–20-fold), RUNX3 (3–15-fold), and Myb (2-fold), and that mutation of each PU.1-binding element did not reduce RUNX-mediated transactivation (Fig. 5A; data not shown). However, the influence of the three factors was considerably weaker in NIH-3T3 cells, suggesting a cell type-specific effect (Fig. 5, B and C). Therefore, PU.1 was transiently cotransfected with either Myb or RUNX3 in NIH-3T3 cells to evaluate their combined effect on the DC-SIGN promoter. Co-expression of PU.1 with either RUNX3 or c-Myb considerably enhanced the activity of the DC-SIGN promoter (Fig. 5, B and C). Moreover, the activity of the DC-SIGN promoter in the presence of PU.1 and either c-Myb or RUNX3 was higher than the activity exhibited in the presence of each individual factor (Fig. 5, B and C). These results demonstrate that the transactivation ability of PU.1 on the DC-SIGN promoter is enhanced in the presence of either Myb or RUNX3 and indicate that PU.1 synergizes with Myb or RUNX3 in transactivation of the DC-SIGN promoter. Interestingly, mutation of the distal PU.1-binding site had a minor effect on the PU.1/RUNX3 or PU.1/Myb collaboration, whereas mutation of the PU.1 element at –77 resulted in an almost complete loss of the collaborative effect (Fig. 5, B and C). Therefore, it can be concluded that the PU.1-binding sites within the DC-SIGN promoter mediate the transactivation ability of PU.1 and that the proximal PU.1-binding site at –77 mediates most of its responsiveness to the collaborative action of PU.1 and either c-Myb or RUNX3.

PU.1 Expression Correlates with DC-SIGN Cell Surface Expression in Monocyte-derived Macrophages and Dendritic Cells and Influences DC-SIGN Expression in THP-1 Cells—The dependence of the DC-SIGN promoter functional activity on the occupancy of the two PU.1-binding sites led us to explore whether a correlation existed between the expression of PU.1 and that of DC-SIGN. Analysis of PU.1 expression in MDDC from two independent donors, where DC-SIGN expression is most prominent (Fig. 6A), indicated the presence of high levels of PU.1 in the nuclei of dendritic cells (Fig. 6B) and revealed that down-regulation of DC-SIGN during MDDC maturation (Fig. 6A) coincided with a considerably decrease in the nuclear content of PU.1 (Fig. 6B). The correlation between the expression of DC-SIGN and PU.1 was not only observed in MDDC maturation but also upon macrophage activation, since nuclear PU.1 was considerably higher in AAM{Phi}, for which DC-SIGN is a cell surface marker (10), than in CAM{Phi} (Fig. 6C). Finally, to determine whether DC-SIGN and PU.1 RNA levels also correlated during dendritic cell maturation, microarray gene profiling experiments were performed on immature and mature MDDC. In agreement with their diminished protein expression in mature MDDC, DC-SIGN and PU.1 RNA were down-regulated to a similar extent during MDDC maturation (between 5 and 8 times in two independent donors, with p < 10–4 for PU.1 mRNA and p < 0.006 for DC-SIGN mRNA), whereas the mRNA for the maturation marker CD83 was greatly increased (p < 10–6)(Fig. 6D). Therefore, and in agreement with the functional relevance of its occupancy of the DC-SIGN promoter, PU.1 expression parallels that of DC-SIGN in activating macrophages and maturing dendritic cells, further suggesting that PU.1 critically contributes to DC-SIGN promoter activity and cell surface expression.



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  		FIGURE 4.
The basal and cell-specific activity of the DC-SIGN promoter is dependent on the –111/–108 and –77/–74 PU.1-binding elements. A, THP-1, K562, and NIH-3T3 cells were transfected with the indicated reporter plasmids, and luciferase activity was determined after 24 h. Promoter activity is expressed relative to the activity produced by the wild-type pCD209–468 reporter plasmid in each transfected cell type after normalization for transfection efficiency. Each experiment was performed three times with two distinct DNA preparations, and a representative experiment is shown. B, THP-1 were transfected with the indicated reporter plasmids, and luciferase activity was determined after 24 h. Promoter activity is expressed relative to the activity produced by the wild-type pCD209–468 reporter plasmid after normalization for transfection efficiency. Data represent mean ± S.D. of triplicate determinations with two different DNA preparations (p < 10–5 for pCD209–468–111MUT and pCD209–468–111/–77MUT and p < 0.05 for pCD209–468–77MUT, when compared with the activity of the wild-type construct). C, NIH-3T3 cells were transfected with the indicated reporter plasmids together with 50 ng of either CMV-0 (empty) or pCDNA3-PU.1 expression plasmids. -Fold induction represents the luciferase activity produced by the pCDNA3-PU.1 expression vector relative to the activity produced by CMV-0. Data represent mean ± S.D. of three independent experiments using two different DNA preparations.

 



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  		FIGURE 5.
The proximal PU.1-binding site within the DC-SIGN gene promoter confers responsiveness to RUNX and c-Myb. A, K562 cells were transfected with the indicated reporter plasmids in the presence of CMV-0, CMV-RUNX1, or CMV-RUNX3, and luciferase activity was determined after 24 h. For each individual reporter construct, -fold induction represents the luciferase activity yielded by an expression vector relative to the activity produced by a similar amount of empty CMV-0 plasmid. Data represent mean ± S.D. of three independent experiments using two different DNA preparations. B and C, NIH-3T3 cells were transfected with the indicated reporter plasmids and either pCDNA3-PU.1 or CMV-RUNX3 + pCDM8-CBF{beta} (B) or CMV-Myb (C) or pCDNA3-PU.1 together with either CMV-RUNX3 + pCDM8-CBF{beta} (B) or CMV-Myb (C). In all cases, total DNA was kept constant (1.5 µg) by adding CMV-0 plasmid DNA, and luciferase activity was determined after 24 h. For each individual reporter construct, -fold induction represents the luciferase activity yielded by an expression vector combination relative to the activity produced by a similar amount of empty CMV-0 plasmid. Data represent mean ± S.D. of three independent experiments using two different DNA preparations.

 
The comparison of AAM{Phi} and CAM{Phi} led us to test whether IL-4 was responsible for the differential expression of PU.1 in both cell types. To that end, monocytes and monocyte-derived macrophages (generated with either granulocyte-macrophage colony-stimulating factor or macrophage colony-stimulating factor) were treated with IL-4, and the expression levels of PU.1 were determined by Western blot. As shown in Fig. 7, PU.1 expression was considerable increased in both monocytes (Fig. 7A) and monocyte-derived macrophages (Fig. 7B), regardless of the cytokine used for macrophage differentiation. As expected, DC-SIGN expression was higher in IL-4-treated monocytes and macrophages, which exhibit higher levels of PU.1 (Fig. 7, A and B). This set of experiments further confirms the correlation between PU.1 and DC-SIGN expression in myeloid cells and indicates that PU.1 expression is regulated by IL-4 in both monocytes and macrophages.



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  		FIGURE 6.
Correlation between PU.1 expression and DC-SIGN cell surface levels in dendritic cells and macrophages. A, cell surface expression of DC-SIGN on immature and lipopolysaccharide-mature monocyte-derived dendritic cells. Flow cytometry was performed using the MR-1 anti-DC-SIGN monoclonal antibody (4) and the supernatant of the murine myeloma P3X63Ag8 as control. B, whole cell extracts were obtained from THP-1, K562, immature (I), and mature (M) MDDC from two independent donors and untransfected or pCDNA3-PU.1-transfected COS-7 cells, and 10 µg of each extract was subjected to Western blot using a polyclonal antiserum specific for human PU.1 (sc-352; Santa Cruz Biotechnology). C, whole cell extracts were obtained from MDDC (immature (I) and mature (M)) and MDM treated for 48 h with either IL-4 (AAMØ) or interferon-{gamma} (CAMØ), and 10 µg of each extract was subjected to Western blot using a polyclonal antiserum specific for human PU.1, DC-SIGN (10), or {beta}-actin as control. D, determination of the mRNA level for PU.1 and DC-SIGN in immature and mature MDDC from two independent donors using the human whole genome DNA chip of the Codelink microarray platform. Data shown represent the mature MDDC/immature MDDC mRNA ratio for PU.1 and DC-SIGN, in a log2 scale. The CD83 mRNA -fold change is also represented as control for MDDC maturation.

 
Finally, to definitively establish the influence of PU.1 on DC-SIGN cell surface expression, DC-SIGN RNA and protein levels were determined in THP-1 cells nucleofected with a PU-1-specific siRNA. Transfection of PU-siRNA in THP-1 cells led to a 70% reduction in the levels of PU.1 protein, as determined by Western blot on whole cell extracts (Fig. 7C). More importantly, reverse transcription-PCR and Western blot revealed that DC-SIGN RNA and protein levels were significantly lower in the PU.1-siRNA-transfected cells than in cells transfected with a control siRNA (Fig. 7, C and D). Therefore, diminishing PU.1 expression has a direct impact on DC-SIGN RNA and protein levels, confirming the relevance of PU.1 for DC-SIGN gene expression.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
DC-SIGN is a cell surface molecule involved in intercellular adhesion and antigen uptake whose function is subverted by numerous pathogens in their strategy to evade immunosurveillance (1, 39) and whose expression is restricted to myeloid dendritic cells and certain macrophages (6–10). In the present report, we present evidence that the PU.1 transcription factor is involved in the basal and tissue-specific expression of DC-SIGN through occupancy of two DNA elements within the proximal regulatory region of the DC-SIGN gene. The integrity of the proximal PU.1-binding site is not only required for PU.1 responsiveness but also mediates the transactivating effects of Myb and RUNX3 on the DC-SIGN proximal regulatory region. Moreover, the expression of DC-SIGN correlates with PU.1 nuclear levels during dendritic cell maturation and upon alternative and classical macrophage activation, and reduction of PU.1 expression through siRNA results in diminished cellular DC-SIGN level. Altogether, these results demonstrate a critical role for PU.1 in DC-SIGN gene expression.

The involvement of PU.1 in the cell-specific expression of DC-SIGN is in line with the known functions of PU.1 in myeloid cell differentiation. The analysis of PU.1-deficient mice revealed that PU.1 is required for the proper development of myeloid progenitors and for the generation of myeloid-derived (26) and thymic (27) dendritic cells. In addition, the PU.1-interacting factor ICSBP (interferon consensus sequence-binding protein) is critical for both early differentiation and final maturation of dendritic cells (40). In the human system, PU.1 is also implicated in the generation of Langerhans cells, since its expression in myeloid progenitors triggers Langerhans cell development, and factors that inhibit PU.1 function prevent Langerhans cell generation (28). Since DC-SIGN mediates relevant effector functions in dendritic cells (antigen uptake and interaction with endothelial and T cells), it is tempting to speculate that an altered expression of DC-SIGN might contribute to the altered dendritic cell generation seen in the absence of PU.1. However, and despite its relevance for DC-SIGN promoter activity, PU.1 is required but is not sufficient for DC-SIGN expression, since PU.1+ B lymphocytes are devoid of DC-SIGN expression, and monocyte-derived macrophages exhibit moderate levels of PU.1 but low/absent levels of DC-SIGN. In this regard, other myeloid-specific factors, in combination with PU.1, might ultimately determine the cell- and state-specific expression of DC-SIGN. Besides recognition by PU.1, the proximal Ets element also confers RUNX3 responsiveness to the DC-SIGN proximal promoter in a PU.1-dependent manner. The participation of RUNX3 in the regulation of DC-SIGN expression would be in agreement with the ability of RUNX3 to modulate the phenotypic and functional maturation of dendritic cells, since bone marrow-derived murine dendritic cells from RUNX3 –/– exhibit accelerated maturation (41). In any event, given the down-regulation of PU.1 (this paper) and the up-regulation of RUNX3 upon dendritic cell maturation (42), it is tempting to speculate that RUNX3 might negatively regulate DC-SIGN expression indendritic cells, an explanation compatible with the context-dependent transcriptional regulatory activity of RUNX factors (43).



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  		FIGURE 7.
IL-4-regulated expression of PU.1 and DC-SIGN in monocytes and macrophages, and influence of PU.1 on DC-SIGN expression level. A, whole cell extracts were obtained from untreated monocytes (–) or monocytes treated with either IL-4 or -6 for 24 h, and 10 µg of each whole cell extract was subjected to Western blot using a polyclonal antiserum specific for human PU.1 or DC-SIGN (10). B, whole cell extracts were obtained from untreated or IL-4-treated MDM generated with either granulocyte-macrophage colony-stimulating factor or macrophage colony-stimulating factor from two independent donors (MDM#1 and MDM#2). 10 µg of each extract was subjected to Western blot using a polyclonal antiserum specific for human PU.1 or DC-SIGN (10). Whole cell extracts from MDDC were analyzed in parallel as control. C, 2 x 106 THP-1 cells was transfected by nucleofection with 3 µg of siRNA for PU.1 (PU.1) or a control siRNA using the Cell Line Nucleofector Kit V (Amaxa). After 24 h, half of the cells were lysed, and cell lysates were subjected to Western blot using a polyclonal antiserum specific for human PU.1, DC-SIGN (10), or {beta}-actin as control. 10 µg of a lysate from untransfected THP-1 cells (lanes labeled Untreated) was analyzed in parallel to mark the positions of the PU.1, DC-SIGN, and {beta}-actin bands and to serve as a reference for antibody specificity. The experiment was performed twice with similar results, and one of the experiments is shown. D,2 x 106 THP-1 cells was transfected by nucleofection with 3 µg of siRNA for PU.1 (PU.1) or a control siRNA using the Cell Line Nucleofector Kit V (Amaxa). After 24 h, one-fifth of the cells were lysed, and cell lysates were subjected to Western blot using a polyclonal antiserum specific for human PU.1 (upper panel). RNA from the remaining siRNA-transfected cells was subjected to reverse transcription-PCR for the indicated cycles to specifically amplify the whole coding region of the DC-SIGN RNA (lower left panels) or a 417-bp fragment from the glyceraldehyde-3-phosphate dehydrogenase RNA as control. Control PCRs were carried out in the absence of cDNA (lanes labeled with a minus sign). Two independent experiments were performed, and both of them are shown.

 
Reverse transcription-PCR experiments have led to the identification of five murine molecules with homology to DC-SIGN (mDC-SIGN and SIGNR1 to -4) (44). Their degree of identity to human DC-SIGN in the lectin domain ranges from 65 to 70%, but all of them exhibit considerably shorter neck domains weakly homologous to that of human DC-SIGN (44). Based on its genomic location and the high level of mRNA in murine CD11+ dendritic cells, one of the molecules has been termed mDC-SIGN, whose mRNA can be detected also in B cells and even spleen T lymphocytes (44). However, comparison of upstream genomic sequences reveals that the sequence around position –111, whose disruption critically affects the activity of the human DC-SIGN promoter (Fig. 4A), is conserved in the putative regulatory regions of SIGNR1, SIGNR3, and SIGNR4, whereas the proximal PU.1-binding site at –77 is maintained within the SIGNR3 and SIGNR4 genes, and none of them is conserved in the mDC-SIGN gene (data not shown). A more extensive analysis of the pattern of expression of all of the DC-SIGN murine homologues is required to evaluate the functional significance that the conservation or loss of these elements might have.

Together with other C-type lectins on the surface of dendritic cells, DC-SIGN appears as a key player in the initial stages of HIV infection through its ability to capture and transfer virus to T lymphocytes (45). DC-SIGN is found in both peripheral blood dendritic cell precursors and mucosal dendritic cells and is therefore adequately positioned for HIV spread both after sexual transmission and after contamination with blood. The participation of DC-SIGN in HIV infection has prompted studies to identify either DC-SIGN variants or polymorphisms that correlate with reduced or increased risk of HIV-1 infection. Heterozygosity of a DC-SIGN variant affecting the "neck" domain has been proposed to reduce the risk of HIV-1 infection (46), whereas a large study on the European-American population at risk for HIV infection has revealed that a single-nucleotide polymorphism in the DC-SIGN promoter associates with an increased risk for parenteral HIV-1 infection (19). This polymorphism affects a nucleotide included within the –468/–19 region and is located upstream of the PU.1-binding sites identified in the present study. The functional relevance of this polymorphism in terms of promoter activity has not been evaluated (19), and it will be interesting to determine the identity of the factors binding to this DNA element as well as their potential cooperation with PU.1 for DC-SIGN gene transcription.

A large battery of receptors are used by phagocytes to recognize both foreign invaders and endogenously produced molecules (47). These receptors include Toll-like receptors, scavenger receptors, complement and Fc receptors, integrins, and members of the C-type lectin receptor family, most of which have evolved to recognize conserved motifs on pathogens that are not found on higher eukaryotes (47). The down-regulation of PU.1 upon dendritic cell maturation suggests that the factor has a more relevant role in the effector functions displayed by immature dendritic cells, whose more characteristic ability is the binding and uptake of extracellular material for subsequent antigen presentation. In this regard, the participation of PU.1 in DC-SIGN expression is in line with previous reports describing the ability of PU.1 to regulate the transcription of genes encoding receptors involved in pathogen recognition and antigen uptake such as Toll-like receptor 4 (48), the scavenger receptor CD68 (49), the integrins CD11c and CD11b (32, 50), and the macrophage mannose receptor (51, 52). Therefore, the involvement of PU.1 in DC-SIGN expression can be framed within the contribution that PU.1 appears to have to the acquisition of the antigen uptake molecular repertoire in dendritic cells and macrophages.

On the other hand, the higher level of PU.1 in AAM{Phi}, as compared with interferon-{gamma}-treated macrophages, and its up-regulation by IL-4 suggest that PU.1 might be preferentially expressed during Th2 immune responses. The involvement of PU.1 in the IL-4-induced expression of arginase I in macrophages (53) further supports this hypothesis, since elevated arginase I expression is characteristic of AAM{Phi} and is responsible for their lack of NO production and reduced microbicidal function (54). The implication of PU.1 in the IL-4-induced expression of the AAM{Phi}-specific markers DC-SIGN and arginase I is consistent with the relative levels of PU.1 that we have observed in AAM{Phi} and CAM{Phi}. Based on these observations, the involvement of PU.1 in the acquisition of other AAM{Phi}-specific effector functions (promotion of tumor cell growth and wound repair, proangiogenic activity, extracellular matrix deposition) is a matter that deserves further investigation.


   FOOTNOTES
 
* This work was supported by Ministerio de Educación y Ciencia Grant SAF2002-04615-C02-01 and Grant GEN2003-20649-C06-01/NAC and Fundación para la Investigación y Prevención del SIDA en España Grant 36422/03 (to A. L. C.) and Grant GEN2003-20649-C06-06/NAC (to M. A. V.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 Supported by a predoctoral grant from Ministerio de Educación y Ciencia (Spain). Back

2 Supported by an I3P postdoctoral contract through CSIC. Back

3 To whom correspondence should be addressed: Centro de Investigaciones Biológicas, CSIC, Ramiro de Maeztu 9, Madrid 28040, Spain. Tel.: 34-91-8373112 (ext. 4376); Fax: 34-91-5627518; E-mail: acorbi{at}cib.csic.es.

4 The abbreviations used are: DC-SIGN, dendritic cell-specific intercellular adhesion molecule-3-grabbing nonintegrin; DC, dendritic cell; IL, interleukin; MDDC, monocyte-derived dendritic cell(s); HIV, human immunodeficiency virus; MDM, monocyte-derived macrophages; EMSA, electrophoretic mobility shift assay; AAM{Phi}, alternatively activated macrophage(s); CAM{Phi}, classically activated macrophage(s); contig, group of overlapping clones; siRNA, small interfering RNA. Back



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 INTRODUCTION
 EXPERIMENTAL PROCEDURES
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 DISCUSSION
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