Current immunological opinion holds that myeloid dendritic cell (mDC) precursors migrate from the blood to the tissues, where they differentiate into immature dermal- and Langerhans-type dendritic cells (DC). Tissue DC require appropriate signals from pathogens or inflammatory cytokines to mature and migrate to secondary lymphoid tissue. We show that purified blood mDC cultured in vitro with GM-CSF and IL-4, but in the absence of added exogenous maturation stimuli, rapidly differentiate into two maturational and phenotypically distinct populations. The major population resembles immature dermal DC, being positive for CD11b, CD1a, and DC-specific ICAM-3-grabbing nonintegrin. They express moderate levels of MHC class II and low levels of costimulatory molecules. The second population is CD11b−/low and lacks CD1a and DC-specific ICAM-3-grabbing nonintegrin but expresses high levels of MHC class II and costimulatory molecules. Expression of CCR7 on the CD11b−/low population and absence on the CD11b+ cells further supports the view that these cells are mature and immature, respectively. Differentiation into mature and immature populations was not blocked by polymyxin B, an inhibitor of LPS. Neither population labeled for Langerin, E-cadherin, or CCR6 molecules expressed by Langerhans cells. Stimulation of 48-h cultured DC with LPS, CD40L, or poly(I:C) caused little increase in MHC or costimulatory molecule expression in the CD11b−/low DC but caused up-regulated expression in the CD11b+ cells. In HIV-infected individuals, there was a marked decrease in the viability of cultured blood mDC, a failure to differentiate into the two populations described for normal donors, and an impaired ability to stimulate T cell proliferation.

Dendritic cells (DC)3 are the most potent of the professional APCs and are the only cells capable of efficiently stimulating naive T cells (1, 2). They are thus essential for the initiation of primary adaptive cellular immune responses. In the absence of infection or inflammatory stimuli, DC reside in the peripheral tissues in an immature state. They express low levels of the costimulatory molecules CD80 and CD86 and are effective at acquiring and processing Ag but are less adept at stimulating T cells than mature DC. Signals from pathogens or the cellular pathology and trauma associated with infection induce maturation and mobilization of tissue DC to the draining lymph node. More specifically, these events are mediated by pathogen-derived molecules such as LPS, unmethylated CpG motifs in bacterial DNA, and dsRNA that stimulate DC through Toll receptors, TNF-α, and heat shock proteins, released from necrotic tissue (3, 4). Mature DC up-regulate MHC class I and II, CD83, and the costimulatory molecules CD80 and CD86, which signal to the T cells through CD28. These changes are accompanied by the expression of the chemokine receptor CCR7 that mediates homing to the lymph node in response to CCL19 and CCL21 (5).

Tissue DC are represented by Langerhans cells, which lie in the epidermal regions of the skin and genital tract mucosal tissue, and by dermal- and interstitial-type DC in most other body tissues. DC are distributed to the tissues via the blood, where their precursors are recognized by the expression of MHC class II, the β integrin CD11c, and lack of lineage markers associated with other mononuclear cell populations (6). The recent recognition that these blood precursors express CD1c, also called BDCA-1, has facilitated their purification using immunomagnetic bead technology, and cells isolated with these reagents have been shown to express myeloid cell markers including CD11c, CD13, and CD33, hence commonly termed myeloid DC (mDC) (7, 8). mDC are distinct from plasmacytoid DC, which do not express CD11c and migrate directly from the blood to the T cell regions of secondary lymphoid tissue, where they secrete high levels of IFN-α in response to virus (9, 10). Although the CD11c-positive blood mDC are generally considered to include precursors of Langerhans cell DC, there is only one report that provides direct evidence to support this view (11).

mDC constitute only ∼0.5% of the mononuclear cell fraction in human blood. Thus, most studies of human DC have used monocyte-derived DC, which can be generated in large numbers with relative ease by culturing monocytes for 5–7 days in GM-CSF and IL-4 (12, 13). These cells appear to be equivalent to dermal-type tissue DC. There is increasing interest in using peripheral blood DC in vaccine-based clinical trials; however, recent data suggest that there may be important phenotypic and functional differences between blood mDC and monocyte-derived DC (7). These differences may be significant in the design of clinical trials to investigate the potential of this cell population to augment adaptive immune responses. In this study, we show that in the absence of maturational stimuli, blood mDC differentiate into two distinct populations on culture with GM-CSF and IL-4. The major population expresses CD11b and resembles immature dermal or monocyte-derived DC. The second population is CD11b−/low and lacks markers for dermal- or Langerhans-type DC but shows an activated or mature phenotype. We also show that, in HIV-1 infection, a condition associated with significant defects in mDC numbers and function (14, 15, 16, 17, 18, 19), there is reduced viability and impaired differentiation into CD11b+ and CD11b−/low populations. These findings may provide an explanation for the functional defect observed in these cells.

Buffy coats were obtained from the United Kingdom National Blood Transfusion Unit (Colindale, London, U.K.). PBMC were isolated by centrifugation over Ficoll-Histopaque and then separated into a low-density fraction enriched in DC by centrifuging over a 50% Percoll gradient for 30 min at 300 × g. This fraction was labeled with mAbs to CD3, CD20, and CD14 (hybridomas UCHT1, 1F54, and 60bca were obtained from the American Tissue Culture Collection), and labeled cells were depleted with anti-Ig-conjugated magnetic beads (Dynal Biotech). CD11c+ DC were then isolated using anti-CD1c immunomagnetic beads (BDCA-1; Miltenyi Biotec) following the manufacturer’s instructions. To assess purity, isolated cells were labeled with a mixture of PE-conjugated Abs to CD3, CD14, CD16, and CD19; PerCP was conjugated to HLA-DR; and FITC was conjugated to anti-CD11c. DC were identified as cells that expressed DR and CD11c but were negative for PE staining. Isolated DC were >95% pure. Purified mDC were cultured in 96-well round-bottom microtiter plates (105/well) in 200 μl of RPMI 1640 buffered with NaHCO3, containing 10% heat-inactivated endotoxin-tested FCS, 100 IU/ml penicillin, 0.1 mg/ml streptomycin, 2 mM l-glutamine (all from Sigma-Aldrich), 100 ng/ml GM-CSF, and 100 U/ml IL-4 (cytokines from R&D Systems). After 48 h, cells were analyzed by FACSCalibur or sorted into CD11b+ and CD11b fractions for T cell proliferation assays. Culture supernatants were assayed for LPS using the Limulus assay (Associates of Cape Cod), following the procedure advocated by the manufacturer.

Ethical approval and informed patient consent was obtained for 50 ml of blood provided by 19 antiretroviral-naive patients. mDC were isolated from PBMC using the BDCA-1 isolation kit (Miltenyi Biotec) as described above. Patient information including CD4 T cell counts and virus load are given in Table I.

Table I.

Summary of patient clinical data

Patient No.AgeSexaCD4 No./μlVirus RNA Copies/ml
49 208 124,959 
38 278 480,000 
27 284 162,745 
40 248 295,346 
23 335 283,523 
31 646 3,378 
44 585 10 
31 313 295,360 
23 162 122,954 
10 42 354 4,750 
11 32 197 364,917 
12 77 382 769 
13 52 372 57,041 
14 28 240 25,210 
15 40 488 1,352 
16 50 314 215,834 
17 32 1022 <50 
18 30 305 95,611 
19 51 14 250,053 
Patient No.AgeSexaCD4 No./μlVirus RNA Copies/ml
49 208 124,959 
38 278 480,000 
27 284 162,745 
40 248 295,346 
23 335 283,523 
31 646 3,378 
44 585 10 
31 313 295,360 
23 162 122,954 
10 42 354 4,750 
11 32 197 364,917 
12 77 382 769 
13 52 372 57,041 
14 28 240 25,210 
15 40 488 1,352 
16 50 314 215,834 
17 32 1022 <50 
18 30 305 95,611 
19 51 14 250,053 
a

M, Male; F, Female.

For the phenotypic characterization of DC by FACSCalibur, the following Abs were obtained from BD Pharmingen: PE conjugated to anti-CD3, -CD14, -CD16, -CD19, and -CD11b; PerCP conjugated to anti-HLA-DR; FITC conjugated to anti-CD80, -CD83, -CD86, -CD1a, and DC-specific ICAM-3-grabbing nonintegrin (DC-SIGN); and allophycocyanin conjugated to anti-CD11c, -CD1a, -CCR5, and -CXCR4. Abs from other sources included FITC anti-CD11b (Serotec), FITC anti-CCR7 (R&D Systems), PE anti-Langerin (Immunotech), and FITC anti-CD11c (DakoCytomation).

Purified mDC were cultured for 48 h with GM-CSF and IL-4 and then stimulated for 24 h with either 1 μg/ml LPS (Sigma-Aldrich), 20 μg/ml poly(I:C) or at a ratio of 1:1 with an irradiated murine myeloma cell line transfected with human CD154/CD40L molecule (20).

Purified myeloid blood DC were cultured for 2 days in GM-CSF and IL-4 and then labeled with PE anti-CD11b and CyChrome-conjugated anti-HLA-DR. The cells were then sorted into CD11b+ and CD11b fractions by using a FACSVantage SE high-speed cell sorter (BD Biosciences). The purity of sorted populations was >98%.

Responder cells for MLRs were prepared from buffy coats by centrifuging PBMC over a 50% Percoll gradient as described above. Pellet cells enriched for T cells and depleted of DC and monocytes were cryopreserved until required as responders in MLRs. Proliferation assays were performed in 96-well round-bottom microtiter plates in RPMI 1640 NaHCO3, containing 100 IU/ml penicillin, 0.1 mg/ml streptomycin, 2 mM l-glutamine, and 10% male AB plasma (all from Sigma-Aldrich). Triplicate doubling dilutions of CD11b+ or CD11b DC were cocultured with 5 × 104 allogeneic responder cells for 6 days. On day 5, the cells were pulsed with 50 μl (0.5 μCi) of tritiated thymidine. Incorporation that occurred during the following 16 h was assessed by a beta plate reader.

Purified blood mDC were maintained for 48 h in GM-CSF and IL-4 and then cultured with FITC-conjugated dextran (Sigma-Aldrich) (molecular mass, 40,500) at 1 mg/ml for 30 min at 37°C. Subsequently, they were labeled on ice with PE-conjugated anti-CD11b and PerCP anti-MHC class II, washed, fixed in 4% paraformaldehyde, and analyzed by flow cytometry. DC were also cultured with FITC-dextran on ice to serve as negative controls for uptake.

After culture for 48 h with cytokines, myeloid blood DC were sorted into CD11b+ and CD11b−/low populations as described above, pulsed with tetanus toxoid Ag (10 μg/ml) for 1 h at 37°C, and then washed. Responder autologous lymphocytes were prepared by depleting Percoll pellet fractions of PBMC of non-DC APCs and enriched for CD4 T cells using magnetic beads conjugated to anti-CD14, anti-CD19, and anti-CD8 (Dynal Biotech). A total of 5 × 104 responder cells (70–80%; CD4) T cells was cocultured in triplicate in 96-well plates with either 5000 or 2500 Ag-pulsed CD11b+ or CD11b−/low DC. Responder cells were also cultured alone, with Ag, or with DC that were not pulsed with Ag.

mDC were isolated from buffy coats of donors, seronegative for HIV, hepatitis B, and hepatitis C, based on the selection of cells expressing the Ag BDCA1 (CD1c) (21). Most buffy coats yielded between 1 and 2 × 106 mDC. These cells were positive for CD11c and expressed high levels of HLA-DR (Fig. 1,a). Analysis of CD11b expression showed a continuum of staining from cells that were totally negative, usually constituting the majority of the population, to cells expressing low and moderate levels of labeling. Gating on cells that were CD11b+ or CD11b, further analysis showed that both populations expressed CD4 and low levels of CD86 and CD40. CD1a, CD80, CD83, and DC-SIGN were not detectable on either population. CD45RO was expressed at a marginally higher level on the CD11b+ DC (Fig. 2,a). Isolated DC cultured in GM-CSF and IL-4, but in the absence of known DC-activating stimuli, differentiated into a major population that expressed CD11b and moderate levels of DR and a second population that was CD11b−/low but expressed high levels of DR (Fig. 1). The number of cells expressing CD11b increased from ≤50% before culture to between 70 and 90% after 72 h. The level of CD11b expression reached maximum levels by 48 h. Differentiation into these two populations was observed consistently in >20 experiments. At 48 h, the median percentage of CD11b−/low DC was 27.2 (n = 29) with 25th and 75th percentiles at 18 and 36.9%, respectively (Fig. 1 g). We also cultured DC in 10% human AB serum instead of FCS and still observed differentiation into the two DC populations.

FIGURE 1.

Differentiation of CD11c-positive blood DC into two populations. mDC were isolated from buffy coats using anti-CD1c conjugated to magnetic beads. a and b, Freshly isolated cells expressed DR and CD11c (a), and a major fraction expressed low to intermediate levels of CD11b (b). c–e, On culture in GM-CSF and IL-4, the cells differentiated into distinct CD11b-positive and -negative or low populations. Differentiation into two populations was observed in experiments on >20 donors. g, The percentage and median of the CD11b−/low DC population at 48 h from 29 separate experiments is shown.

FIGURE 1.

Differentiation of CD11c-positive blood DC into two populations. mDC were isolated from buffy coats using anti-CD1c conjugated to magnetic beads. a and b, Freshly isolated cells expressed DR and CD11c (a), and a major fraction expressed low to intermediate levels of CD11b (b). c–e, On culture in GM-CSF and IL-4, the cells differentiated into distinct CD11b-positive and -negative or low populations. Differentiation into two populations was observed in experiments on >20 donors. g, The percentage and median of the CD11b−/low DC population at 48 h from 29 separate experiments is shown.

Close modal
FIGURE 2.

Maturation of DC in the absence of stimulation. a, Phenotype of human mDC immediately after isolation from blood or after 48-h culture in GM-CSF and IL-4. Dividing freshly isolated cells into CD11b+ and CD11b revealed that both populations expressed similar levels of costimulatory molecules and CD4, but neither population expressed CD1a or DC-SIGN. CD45RO was present at higher levels on the CD11b+ population. b, After 48-h culture, the CD11b+ DC became positive for CD1a and DC-SIGN, whereas the CD11b DC lacked these surface markers but expressed CD83 and CD80 and showed higher levels of CD86. There was a more marked down-regulation of CD4 in the CD11b+ population. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. Each FACSCalibur profile is representative of results obtained from experiments performed on DC from a minimum of four donors.

FIGURE 2.

Maturation of DC in the absence of stimulation. a, Phenotype of human mDC immediately after isolation from blood or after 48-h culture in GM-CSF and IL-4. Dividing freshly isolated cells into CD11b+ and CD11b revealed that both populations expressed similar levels of costimulatory molecules and CD4, but neither population expressed CD1a or DC-SIGN. CD45RO was present at higher levels on the CD11b+ population. b, After 48-h culture, the CD11b+ DC became positive for CD1a and DC-SIGN, whereas the CD11b DC lacked these surface markers but expressed CD83 and CD80 and showed higher levels of CD86. There was a more marked down-regulation of CD4 in the CD11b+ population. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. Each FACSCalibur profile is representative of results obtained from experiments performed on DC from a minimum of four donors.

Close modal

The consensus view is that blood mDC give rise to tissue dermal- and Langerhans-type DC. To provide evidence in support of this hypothesis, we performed a detailed phenotypical analysis of cultured blood CD11c+ DC. At 48 h, the CD11b+ DC population expressed CD1a, a molecule found on dermal- and Langerhans-type DC and the C-type lectin, DC-SIGN, a molecule uniquely found on dermal and monocyte-derived DC. Neither of these molecules was detected on the CD11b−/low DC (Fig. 2,b). The two DC populations were also distinct with respect to their maturational status. On encountering inflammatory cytokines or components of microorganisms that are recognized by TLRs, DC show up-regulation of MHC and costimulatory molecules. Despite culture in the absence of exogenously added maturational stimuli, the CD11b−/low DC showed a mature phenotype. By contrast, CD11b+ DC in the same cultures showed an immature phenotype. The CD11b−/low DC consistently expressed higher levels of MHC class II, CD80, and CD86. CD83, a molecule expressed on mature but not by immature DC, was always observed on CD11b−/low DC in unstimulated cultures but never detected on CD11b+ DC. LPS was not detected in 48-h culture supernatant by the Limulus assay, but to further rule out the possibility that maturation of CD11b−/low DC was mediated by low levels of contaminating endotoxin, cultures were performed in the presence of polymyxin B (1 μg/ml), an inhibitor of LPS-mediated DC activation. Polymyxin B failed to block differentiation into two cell populations or alter expression of MHC and costimulatory molecules on either DC population (results not shown). Because tissue dermal-type DC, but not Langerhans cells, express CD11b and DC-SIGN (22), we speculated that the two populations of DC may represent the precursors of dermal- and Langerhans-type DC. However, we failed to detect the Langerhans cell-associated markers, Langerin, E-cadherin, or CCR6 in freshly isolated or cultured cells, even when TGF-β (1 ng/ml), a cytokine that promotes Langerhans cell differentiation (23), was included in the culture medium (data not shown). Thus, BDCA-1+ blood DC cultured in the absence of exogenous stimuli differentiate into the following two populations: 1) a mature population that is negative for CD11b and DC-SIGN but expresses high levels of MHC class II, CD80, CD83, and CD86; and 2) an immature CD11b/DC-SIGN+ population that expresses lower levels MHC class 11 and CD86 and is negative for CD80 and CD83. For comparison, we generated DC from monocytes isolated from Percoll interface cells using magnetic beads conjugated to anti-CD14 and culturing for 6 days in GM-CSF and IL-4. Monocyte-derived DC expressed CD11b, CD1a, DC-SIGN, and moderate levels of CD80 and CD86, but were negative for CD83 (Fig. 3). There was no evidence of differentiation into two phenotypically distinct populations, as was observed with the cultured blood mDC.

FIGURE 3.

The phenotypic profile of DC generated by culturing monocytes with GM-CSF and IL-4 for 6 days. Immature monocyte-derived DC uniformly express CD11b and are positive for CD1a, and DC-SIGN express moderate levels of CD80 and CD86 but do not express CD83.

FIGURE 3.

The phenotypic profile of DC generated by culturing monocytes with GM-CSF and IL-4 for 6 days. Immature monocyte-derived DC uniformly express CD11b and are positive for CD1a, and DC-SIGN express moderate levels of CD80 and CD86 but do not express CD83.

Close modal

The functional hallmark of a DC is the ability to stimulate T cell proliferation. We therefore sorted the DC after 48-h culture into CD11b+ and CD11b DC and measured their ability to stimulate T cell proliferation in an allogeneic MLR. In five independent experiments, both populations effectively stimulated allogeneic T cell proliferation (Fig. 4 a).

FIGURE 4.

Functional properties of CD11b+ and CD11b DC. a, Purified blood mDC cultured for 48 h in GM-CSF and IL-4 and sorted into CD11b+ and CD11b fractions. DC populations were titrated into a 96-well plate, and 5 × 104 allogeneic lymphocytes, separated from DC and monocytes by centrifugation over a 50% Percoll gradient, were added to each well. □, Stimulation by CD11b DC. ⋄, Stimulation by CD11b+ DC. Bars indicate SD; the negative bar has been omitted for clarity. After 5 days, cells were pulsed with [3H]thymidine and harvested after 16 h. Background counts of lymphocytes alone were <50. These results are representative of data obtained in experiments on DC from five donors. b, CD11b+ DC show high levels of FITC-dextran uptake, whereas uptake by CD11b DC was low or undetectable. Overnight stimulation with LPS induced maturation of the CD11b+, as evidenced by increased DR expression, and reduced the ability to take up FITC dextran. Filled histograms represent FITC uptake in cells incubated on ice, and open histograms show uptake in cells incubated at 37°C. One of four representative experiments is shown. c, Both populations of DC were shown to be able to present tetanus toxoid to autologous CD4 T cells. Purified blood mDC were cultured for 48 h in GM-CSF and IL-4, sorted into CD11b+ (DC+) and CD11b−/low (DC) fractions, and cultured for 1 h with tetanus toxoid. After washing, Ag-pulsed DC were cocultured with 50,000 CD4-enriched T cells for 6 days and pulsed with [3H]thymidine for the last 18 h of culture. T cells were also cultured alone, with Ag, or with DC that had not been pulsed with Ag. All assays were performed in triplicate; error bars indicate SD. Similar results were obtained in two separate experiments.

FIGURE 4.

Functional properties of CD11b+ and CD11b DC. a, Purified blood mDC cultured for 48 h in GM-CSF and IL-4 and sorted into CD11b+ and CD11b fractions. DC populations were titrated into a 96-well plate, and 5 × 104 allogeneic lymphocytes, separated from DC and monocytes by centrifugation over a 50% Percoll gradient, were added to each well. □, Stimulation by CD11b DC. ⋄, Stimulation by CD11b+ DC. Bars indicate SD; the negative bar has been omitted for clarity. After 5 days, cells were pulsed with [3H]thymidine and harvested after 16 h. Background counts of lymphocytes alone were <50. These results are representative of data obtained in experiments on DC from five donors. b, CD11b+ DC show high levels of FITC-dextran uptake, whereas uptake by CD11b DC was low or undetectable. Overnight stimulation with LPS induced maturation of the CD11b+, as evidenced by increased DR expression, and reduced the ability to take up FITC dextran. Filled histograms represent FITC uptake in cells incubated on ice, and open histograms show uptake in cells incubated at 37°C. One of four representative experiments is shown. c, Both populations of DC were shown to be able to present tetanus toxoid to autologous CD4 T cells. Purified blood mDC were cultured for 48 h in GM-CSF and IL-4, sorted into CD11b+ (DC+) and CD11b−/low (DC) fractions, and cultured for 1 h with tetanus toxoid. After washing, Ag-pulsed DC were cocultured with 50,000 CD4-enriched T cells for 6 days and pulsed with [3H]thymidine for the last 18 h of culture. T cells were also cultured alone, with Ag, or with DC that had not been pulsed with Ag. All assays were performed in triplicate; error bars indicate SD. Similar results were obtained in two separate experiments.

Close modal

As DC mature, their endocytic capacity is down-regulated. We therefore compared the ability of the two DC populations to endocytose a FITC-dextran conjugate. In four separate experiments, we observed low or undetectable uptake of FITC-dextran by the CD11b−/low DC and high levels of endocytosis by the CD11b+ population (Fig. 4 b). Stimulation for 18 h with LPS up-regulated MHC class II expression of the CD11b+ DC but reduced their endocytic capacity.

Because the CD11b+ DC showed higher levels of endocytosis, we wondered whether there were also differences in their ability to internalize, process, and present an exogenous Ag. Sorted CD11b+ and CD11b−/low DC were pulsed with tetanus toxoid for 1 h, washed, and cocultured with autologous lymphocytes depleted of monocytes, B cells, and CD8 T cells. Both populations of tetanus toxoid-pulsed DC augmented T cell proliferation, although the difference in the level of T cell proliferation between cultures containing unpulsed and pulsed DC was greater for the CD11b+ DC (Fig. 4 c). The higher levels of autologous T cell stimulation observed with the unpulsed CD11b−/low DC may reflect higher expression of DR and costimulatory molecules. Processing and presentation of tetanus toxoid by both DC populations was observed in assays using cells from two different donors.

IL-4 is required for the in vitro generation of DC from monocytes. We therefore determined whether IL-4 was required for the differentiation of DC into the two populations. Myeloid blood DC cultured with GM-CSF alone differentiated into CD11b+ and CD11b−/low DC with no marked difference in CD1a or CD83 expression. However, in the absence of IL-4, there was only weak expression of DC-SIGN on the CD11b+ population (Fig. 5). Because the CD11b−/low cells displayed a more mature phenotype after 48-h culture, we questioned whether CD11b+ cells are the precursors of the CD11b−/low DC. To test this hypothesis, 48-h cultured cells were stimulated for a further 24 h with LPS, poly(I:C), or irradiated CD40L-transfected mouse myeloma cells. Distinct CD11b+ and CD11b−/low DC populations could still be identified in stimulated cultures after 24 h (Fig. 6,a). However, as expected, there was marked up-regulation of MHC class II, CD80, CD83, and CD86 molecules on the CD11b+ population (Fig. 6,b). Marginal or no changes were observed in the expression of these markers on the CD11b−/low DC, suggesting that they matured fully in the absence of added exogenous stimuli. There was a slight reduction in the expression of CD1a and DC-SIGN on stimulated CD11b+ DC. These findings suggest that the CD11b+ DC do not readily differentiate into cells with CD11b−/low phenotype. To provide further evidence that the CD11b+ DC do not differentiate into CD11b−/low cells or vice versa, cells were sorted after 48 h and then cultured in GM-CSF and IL-4 for a further 20 h. Sorted cultured cells showed no change in CD11b expression, thus supporting the view that they represent stable phenotypes that do not readily interconvert (Fig. 7). Interestingly, sorted cultured CD11b+ DC maintained higher viability than the CD11b population. In two experiments, the viability of the CD11b+CD11b DC was 81 vs 37% and 76 vs 50%. This observation could explain the gradual increase in percentage of CD11b+ cells in the mixed cultures (Fig. 1).

FIGURE 5.

IL-4 enhances DC-SIGN expression. Purified blood DC were cultured with GM-CSF and IL-4 or with GM-CSF alone. IL-4 was not required for differentiation into CD11b-positive cells expressing CD1a- and CD11b-negative populations, but, in the absence of IL-4, there was only low-level expression of DC-SIGN on the CD11b+ DC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from four donors.

FIGURE 5.

IL-4 enhances DC-SIGN expression. Purified blood DC were cultured with GM-CSF and IL-4 or with GM-CSF alone. IL-4 was not required for differentiation into CD11b-positive cells expressing CD1a- and CD11b-negative populations, but, in the absence of IL-4, there was only low-level expression of DC-SIGN on the CD11b+ DC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from four donors.

Close modal
FIGURE 6.

The effect of maturation stimuli on 48-h cultures of blood DC. a, Cultures stimulated for 24 h with LPS, poly(I:C), or CD40L maintained distinct CD11b+ and CD11b populations. b, All the maturational stimuli caused up-regulation of DR, CD80, CD83, and CD86 on the CD11b+ cells but caused minimal or no changes in the level of expression of these molecules on the CD11b−/low population. Stimulation resulted in the slight reduction in expression of CD1a and DC-SIGN on the CD11b+ DC. In all plots, the filled histograms represent staining of unstimulated control cells, whereas open histograms show labeling of cells in stimulated cultures. These results are representative of experiments performed on DC from four donors.

FIGURE 6.

The effect of maturation stimuli on 48-h cultures of blood DC. a, Cultures stimulated for 24 h with LPS, poly(I:C), or CD40L maintained distinct CD11b+ and CD11b populations. b, All the maturational stimuli caused up-regulation of DR, CD80, CD83, and CD86 on the CD11b+ cells but caused minimal or no changes in the level of expression of these molecules on the CD11b−/low population. Stimulation resulted in the slight reduction in expression of CD1a and DC-SIGN on the CD11b+ DC. In all plots, the filled histograms represent staining of unstimulated control cells, whereas open histograms show labeling of cells in stimulated cultures. These results are representative of experiments performed on DC from four donors.

Close modal
FIGURE 7.

Cultured DC maintain their CD11b phenotype. To confirm that CD11b+ DC do not readily differentiate into CD11b cells, or vice versa, 48-h cultures (a) were sorted into CD11b+ and CD11b fractions and cultured for a further 20 h. The two populations remained unchanged with respect to CD11b expression (b and c). Identical results were obtained in experiments on two different donors.

FIGURE 7.

Cultured DC maintain their CD11b phenotype. To confirm that CD11b+ DC do not readily differentiate into CD11b cells, or vice versa, 48-h cultures (a) were sorted into CD11b+ and CD11b fractions and cultured for a further 20 h. The two populations remained unchanged with respect to CD11b expression (b and c). Identical results were obtained in experiments on two different donors.

Close modal

It is well established that DC can be derived from monocytes by culturing in the presence of GM-CSF and IL-4, and it is also known that the dermal-type DC derived from CD34 stem cell precursors pass through a CD14 stage during their differentiation (24). Because the phenotypic profile of the CD11b+ population of DC closely resembled that of monocyte-derived DC, we looked for the expression of CD14 on CD1c+BDCA-1+, CD19-negative cells in uncultured blood mononuclear cells (Fig. 8). Fig. 8,a shows that some blood CD1c+BDCA-1+ cells also express CD14 but at a lower level than that on blood monocytes. Analysis of freshly purified DC showed that CD14 expression was predominantly on the CD11b+ population (Fig. 8 b). After 48 h, culture in GM-CSF and IL-4 CD14 became undetectable (not shown).

FIGURE 8.

Uncultured CD11b+ DC express CD14. a, mDC were identified in a PBMC preparation as cells that were negative for CD19 but expressed CD1c (BDCA-1), and a minor fraction was found to express CD14. Expression of CD14 on monocytes is shown for comparison. b, CD14 labeling is shown to be restricted to the CD11b+ fraction of freshly isolated mDC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from four donors.

FIGURE 8.

Uncultured CD11b+ DC express CD14. a, mDC were identified in a PBMC preparation as cells that were negative for CD19 but expressed CD1c (BDCA-1), and a minor fraction was found to express CD14. Expression of CD14 on monocytes is shown for comparison. b, CD14 labeling is shown to be restricted to the CD11b+ fraction of freshly isolated mDC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from four donors.

Close modal

It is well established that chemokine receptor expression on DC is modulated during maturation to facilitate migration to lymphoid tissue. We therefore analyzed expression of CCR7, CXCR4, and CCR5 on freshly isolated DC and DC cultured for 48 h in GM-CSF and IL-4 (Fig. 9). Both populations of freshly isolated DC expressed CXCR4 and low levels of CCR5 but were negative for CCR7 (Fig. 9,a). After 48 h in culture, CCR7 appeared on the surface of the CD11b−/low population and was either absent or present at barely detectable levels on the CD11b+ DC (Fig. 9 b). The levels of CCR5 and CXCR4 on cultured DC showed more experiment-to-experiment variation. However, CXCR4 was expressed consistently at higher levels than CCR5 on both populations and at higher levels on the CD11b−/low DC.

FIGURE 9.

Cultured unstimulated CD11b-negative DC express CCR7. a, Freshly isolated CD11b+ and CD11b−/low DC express CXCR4 and low levels of CCR5 but do not express CCR7. b, After 48-h culture, CCR7 is expressed on CD11b−/low but not CD11b+-positive DC. There was experiment-to-experiment variation in levels of CCR5 and CXCR4 expression on cultured DC, but CXCR4 was expressed consistently at higher levels on the CD11b−/low DC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from a minimum of four donors.

FIGURE 9.

Cultured unstimulated CD11b-negative DC express CCR7. a, Freshly isolated CD11b+ and CD11b−/low DC express CXCR4 and low levels of CCR5 but do not express CCR7. b, After 48-h culture, CCR7 is expressed on CD11b−/low but not CD11b+-positive DC. There was experiment-to-experiment variation in levels of CCR5 and CXCR4 expression on cultured DC, but CXCR4 was expressed consistently at higher levels on the CD11b−/low DC. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. These results are representative of experiments performed on DC from a minimum of four donors.

Close modal

In a recent investigation, we found that blood DC from HIV-infected individuals were impaired in their ability to stimulate T cell proliferation but were unable to identify the mechanisms involved (19). In view of the present findings, we analyzed the differentiation of CD11c+ blood DC from antiretroviral drug-naive HIV-infected patients. However, because <105 mDC were isolated routinely from patient samples, it was not possible to perform as comprehensive an analysis on these cells. We therefore focused on the ability of the cells to differentiate into CD11b+CD1a+ and CD11bCD83+ populations. Analysis of the forward- and side-scatter profiles of 19 patients’ DC cultured for 48 h in GM-CSF and IL-4 showed that there was a marked reduction in viability, mean viability of 23% and range of 1–70%, compared with mean viability of 75% and range of 70–80% in healthy individuals. Only five patients’ samples had >50% of cells in the “live gate” (Table II and Fig. 10,a). Poor viability correlated with impaired ability to differentiate into CD11b+CD1a+ and CD11b−/lowCD83+ DC (Fig. 10, b–d, and Table II). Unlike healthy individuals, no HIV-1 patient sample differentiated into a distinct CD11b+-moderate DR expression subset and a CD11b−/low, strongly positive DR subpopulation. In seven patients (patients 5, 7, 8, 10, 17, 18, and 19), DC differentiated into discrete CD11b+ and CD11b−/low populations; however, in all cases, the expression of DR was equivalent in both DC subsets, and, in two of these (patients 7 and 8), CD1a was not expressed by CD11b+ DC (Fig. 10,d). There was no apparent correlation between differentiation into the DC subsets and virus load, CD4 T cell counts, CD8 T cells numbers, or disease stage 6 mo after analysis of DC function. Of the five patients showing normal differentiation, three had a virus load of >90,000 virus RNA copies/ml, and two patients had virus loads of <5,000 virus RNA copies/ml. In some patients (patients 2, 4, 6, and 12), virtually all the cells remained CD11b (Fig. 10,b), whereas in others (patients 1, 3, 9, 11, 13, 14, 15, and 16), there was a relatively low level expression of CD11b+, and they did not differentiate into separate discrete populations (Fig. 10,c). On analysis of CD1a and CD83 expression of these latter patients, some labeled for neither marker (patients 9 and 11), some labeled for one marker (patients 1 and 15), and three (patients 13, 14, and 16) labeled for both molecules (Table II).

Table II.

Summary of in vitro BDCA-1 DC differentiation in cells isolated from antiretroviral-naive HIV-1-infected patients

Patient No.% Live CellsaCD11bCD1aCD83Max SI
16 Lowb Lowc No NT 
22 No NT No NT 
10 Lowb NT NT NT 
No No No NT 
54 Yes Yes Yes NT 
No No No NT 
29 Yes No Yes NT 
Yes No Yes NT 
Lowb No No NT 
10 70 Yes Yes Yes NT 
11 Lowb No No NT 
12 No No No <2 
13 Lowb Yes Yes <2 
14 16 Lowb Yes Yes <2 
15 Lowb Low No 
16 17 Lowb Low Low 
17 54 Yes Yes Yes 14 
18 55 Yes Yes Yes 
19 58 Yes Yes Yes 
Patient No.% Live CellsaCD11bCD1aCD83Max SI
16 Lowb Lowc No NT 
22 No NT No NT 
10 Lowb NT NT NT 
No No No NT 
54 Yes Yes Yes NT 
No No No NT 
29 Yes No Yes NT 
Yes No Yes NT 
Lowb No No NT 
10 70 Yes Yes Yes NT 
11 Lowb No No NT 
12 No No No <2 
13 Lowb Yes Yes <2 
14 16 Lowb Yes Yes <2 
15 Lowb Low No 
16 17 Lowb Low Low 
17 54 Yes Yes Yes 14 
18 55 Yes Yes Yes 
19 58 Yes Yes Yes 
a

Percentage of cells in live gate from 48-h cultures. Control viability: mean, 75%; range, 70–81%. NT, Not tested. Patient 16 was a recent seroconvertor. Control SI, 51; mean of 3.

b

Cells not differentiated into discrete CD11b-negative and -positive populations.

c

Two pecent of DC expressed CD11b, and these showed low-level CD1a expression.

FIGURE 10.

DC from HIV-infected individuals show impaired differentiation. Myeloid CD1c+ DC were isolated from drug-naive HIV-infected individuals and cultured for 48 h in GM-CSF and IL-4. a, Forward-scatter/side-scatter profiles of one control, patient 11, showing poor DC survival, and patient 10, in which there was normal DC differentiation. b, Patient 2: DC do not differentiate into CD11b+ cells on culture. c, Patient 1 shows cells with low-CD11b expression but minimal expression of CD1a or CD83. d, Patient 7: DC differentiate into CD11b+CD1a and CD11bCD83+ populations. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. (Note: FACSCalibur settings in b are different from those used in c and d.) FACSCalibur analysis data on all 11 patients analyzed are shown in Table II.

FIGURE 10.

DC from HIV-infected individuals show impaired differentiation. Myeloid CD1c+ DC were isolated from drug-naive HIV-infected individuals and cultured for 48 h in GM-CSF and IL-4. a, Forward-scatter/side-scatter profiles of one control, patient 11, showing poor DC survival, and patient 10, in which there was normal DC differentiation. b, Patient 2: DC do not differentiate into CD11b+ cells on culture. c, Patient 1 shows cells with low-CD11b expression but minimal expression of CD1a or CD83. d, Patient 7: DC differentiate into CD11b+CD1a and CD11bCD83+ populations. In all plots, the filled histograms represent staining with an isotype control Ab, whereas open histograms show Ab-specific labeling. (Note: FACSCalibur settings in b are different from those used in c and d.) FACSCalibur analysis data on all 11 patients analyzed are shown in Table II.

Close modal

In view of our initial observations of impaired differentiation, made on patients 1–11, the investigations were extended to analyze T cell stimulatory capacity by MLR. DC were titrated in doubling dilutions, starting with 4000 DC/well, and results were expressed as the highest stimulation index (SI) (3H counts from DC + lymphocytes/3H counts from lymphocytes alone) observed. In parallel, we analyzed the phenotype of these DC after culturing for 48 h. The results of these experiments are summarized in Table II, patients 12–19. All patient DC stimulated T allogeneic T cell proliferation poorly (highest SI, 14) compared with controls (SI, 51; mean of three control donors) and confirm our previous studies (19). An SI >4 was only observed in three patients (patients 16, 17, and 19), and for two of these patients (patients 17 and 19), we observed differentiation into CD11b+ and CD11b−/low populations expressing CD1a and CD83. Of note was that DC from patient 17, showing the highest stimulatory capacity, had an undetectable virus load and a CD4 count of >1000. Surprisingly, the second highest DC stimulatory capacity was seen in patient 19, who had the highest virus load and lowest CD4 number. However, this patient was from Ethiopia, and thus the higher allogeneic T cell stimulation may reflect a greater MHC disparity, and it is also likely that he was infected with a non-clade B virus. For three patients (patients 12, 13, and 14), there were insignificant levels of proliferation. These latter cultures showed low viability; there was no expression of CD11b in one sample (patient 12), whereas the other two expressed low levels of CD11b. DC from patient 18 failed to stimulate a significant MLR despite good cell viability and relatively normal differentiation.

The maturation of DCs into competent APCs that express high levels of MHC and costimulatory molecules is thought to be mediated by inflammatory cytokines or pathogen-derived molecules that are recognized by Toll receptors. Remarkably, in the absence of added maturational stimuli, we observed the differentiation of a small fraction of CD11b−/low, CD1a DC that displayed high-level expression of activation markers CD80, CD83, CD86, high-MHC class II, and CCR7. The absence of detectable LPS, the failure of polymyxin B to inhibit CD11b−/low DC maturation, and the presence of immature CD11b+ DC in the same cultures argues against the presence of exogenous stimulatory molecules. It may be postulated that the CD11b−/low DC population is exquisitely sensitive to low levels of stimulatory signals. However, because one of the main functions of the CD11b+ dermal DC is to serve as a sentinel and rapidly respond to invading pathogens, it is difficult to understand why these latter cells should not be at least equally sensitive to activation stimuli. A more likely explanation is that CD11b−/low DC are programmed to mature in the absence of inflammatory or pathogen-derived signals. Additionally, the expression of CCR7 on these cells suggests that they are programmed to migrate directly to lymphoid tissue. At this stage, we do not know whether factors in the serum are required for differentiation into the two DC populations, and future studies should investigate differentiation in semiartificial medium in addition to the important question of cytokine production by these cells.

The ability of the two DC populations to internalize FITC-dextran conjugates correlated with their maturational status, and thus it was somewhat surprising to find that the mature CD11b−/low population was also able to internalize, process, and present an exogenous Ag. These observations suggest that the uptake of relatively low levels of Ag by the CD11b−/low DC is sufficient to stimulate Ag-specific T cell proliferation. The difference between the level of T cell proliferation in cultures containing Ag-pulsed vs -unpulsed DC was greater for the CD11b+ DC and may reflect the greater endocytic capacity of these cells.

Precursors of tissue DC, Langerhans and dermal DC, are believed to circulate in the blood as a clearly identifiable population of cells that express MHC class II HLA-DR and CD11c, but lack lineage markers associated with T, B, NK, and monocytic cells. In human tissues, both dermal and Langerhans DC express MHC class II and CD1a molecules, but only dermal DC express CD11b (22). These two types of tissue DC can be further differentiated by expression of Birbeck granules, Langerin, and E-cadherin on Langerhans cells and by expression of DC-SIGN on dermal DC. In this study, we show for the first time that CD11c+ DC purified from blood on the basis of BDCA-1 (CD1c) expression rapidly differentiate into two distinct DC populations in culture and that neither population exhibit markers specifically associated with Langerhans cells. The major population identified after 48-h culture was phenotypically identical with immature dermal DC characterized by expression of CD11b, CD1a, DC-SIGN, low levels of CD86, and the absence of membrane staining for the DC maturation markers CD80 and CD83. The lack of CCR7, required for migration to the lymph node in response to CCL19 and CCL21 (5), also supports the conclusion that the CD11b+ population is immature. Although IL-4 was not required for differentiation into CD11b+ and CD11b−/low DC, it did facilitate DC-SIGN expression on the CD11b+ DC. Similarly, there is a requirement for IL-4 for expression of DC-SIGN on monocyte-derived DC (25). Investigations of DC differentiation from CD34+ cord blood stem cells have shown that the precursors of dermal DC express CD14 at an intermediate stage of their development (24). In this study, we show that some BDCA-1+CD11b+ DC that differentiate into dermal DC also express low levels of CD14. Some DC studies have depleted CD11b+ cells before isolating DC, based on reports suggesting that blood DC do not express this marker. However, the present investigations show that a major population of blood-derived DC express CD11b even before culture and may explain the inability to detect DC-SIGN in a previous study (7).

The two DC populations derived from the BDCA-1+CD11c+ blood precursors observed after 48 h are phenotypically distinct, suggesting that they may represent two different differentiation pathways. Both populations were shown to stimulate allogeneic T cell proliferation and thus functionally behave as DC. There was no evidence to support the idea that the immature CD11b+ DC are the precursors of the mature CD11b−/low population. LPS, poly(I:C), or CD40L that promote DC maturation by stimulating through distinct receptors, Toll 4, Toll 3, and CD40, respectively, induced up-regulation of MHC and costimulatory molecules on the dermal-type DC while maintaining expression of CD11b, CD1a, and DC-SIGN. Sorting 48-h cultures into CD11b+ and CD11b cell populations and culturing overnight further confirmed the view that the CD11b+ DC do not readily differentiate into CD11b DC. The appearance of DC-expressing CD83 after short-term culture has been described previously; however, no significance was attached to the fact that this occurred in the absence of activation stimuli (26, 27).

Our work agrees and extends previous observations that multiple subpopulations of CD1c+ peripheral blood DC are likely to exist. Maraskovsky and colleagues (8) have identified distinct CD1c+ peripheral blood DC subsets in the peripheral blood of individuals who had received Flt3 ligand therapy in a trial to DC vaccination in subjects with a history of melanoma. The phenotype of the major DC subset identified in this study after 24 h of culture in GM-CSF and IL-4 was similar to the CD11b−/low that we have seen (CD1a-negative, CD83-positive), and the authors noted that this DC subset had spontaneously up-regulated the expression of CD80, CD83, and DR after overnight culture. In contrast to our findings, they do not describe the cells with an immature phenotype after culture. These differences may be because their studies used blood DC from cancer vaccine patients who had been treated with Flt3 ligand, which may influence the differentiation pathways of CD1c+ DC.

Intriguingly, two distinct populations of CD11c+ DC have been reported in human thymus (28), a minor CD11b+ and a major CD11b population. The former expressed CD64 and moderate levels of CD45RA and was negative for CD83. The latter expressed low CD45RA but high levels of CD83, CD80, CD86, and MHC class II. In these respects, this major thymic DC population is similar to the CD11b−/low cells observed in the present study. It has been suggested that thymic DC differentiate in situ rather than being seeded from the blood (28); however, in view of the present findings, the possibility that some of these DC arise from precursors in the blood should be considered. One difficulty with this hypothesis is that the CD11b−/low blood DC express CCR7, suggesting that they could home to the lymph node. Although immature DC are thought to mediate anergy in the periphery, the observation that the major population in the thymus is mature may indicate that mature DC are required to mediate clonal deletion of newly generated T cells. Under these circumstances, it seems unlikely that thymic CD11b DC depend on inflammatory or pathogenic signals to become activated but may be programmed to develop directly into mature cells. The CD1 family of molecules facilitate the presentation microbial lipids to T cells (29), and recent studies have shown that DC-SIGN can direct exogenous Ag to the MHC class I presentation pathway (30). The absence of these molecules in the CD11b−/low DC population may indicate that the range of Ags that can be presented by these cells is more limited. In the present investigations, we have not analyzed cytokine production, ability to stimulate Th1 or Th2 responses, Ag uptake or processing, and presentation by these mDC populations. Such future studies may help functional elucidation of these cells.

Impaired DC function has been described in a number of human virus diseases including measles, hepatitis C, and HIV (14, 19, 31, 32, 33, 34). In HIV infection, although HIV provirus can be detected in DC, the level of virus detected, <1%, is too low for the functional defect to be directly attributable to infection. The present studies suggest that lack of differentiation may be one factor that contributes to DC dysfunction. The differentiation of BDCA-1+CD11c+ blood DC into two distinct populations was observed consistently in >20 separate experiments on blood obtained from normal uninfected controls. Thus far, we have analyzed the in vitro differentiation of blood DC from 19 antiretroviral drug-naive HIV patients. In 14 of these patients, we observed reduced DC survival and a failure of normal differentiation. Functional analysis in eight samples revealed that all were severely impaired in their ability to stimulate allogeneic T cell proliferation, despite reasonable viability and differentiation into discrete CD11b+ and CD11b−/low populations in three cases. Interestingly, the highest allogeneic stimulatory capacity was observed in patient 17 who showed normal DC differentiation, good DC viability, high CD4 T cell number, and a virus load of <50; nevertheless, the SI value for this patient was still well below that of controls (14 vs 51). These findings suggest that factors additional to viability and differentiation into CD11b+ and CD11b−/low are required for functional competency. Differentiation into the two DC populations was observed in five patients but did not correlate with virus load; three of the five had a virus load >90,000, and two had a virus load of <5,000. To date, the differentiation of blood DC in hepatitis C and measles virus-infected individuals has not been investigated but, in view of the present findings, may be worthwhile.

The underlying mechanism of poor survival has not been elucidated in this study, but to start to address this question, we recently have analyzed expression annexin V on CD11c+ DC from freshly isolated PBMC from anti-retroviral-naive HIV-infected patients (H. Donaghy, P. Kelleher, and S. Patterson, unpublished observations). We found no difference between uninfected control and patient DC in the level of annexin V expression. Survival and differentiation of blood DC in vitro was shown previously to be dependent on GM-CSF (35); thus, future studies on DC from HIV-infected individuals should examine the expression of the GM-CSF receptor.

Investigations in SIV-infected macaques report that, in late-stage disease, there is disruption of DC homeostasis and reduced expression of CD83 and DC-SIGN in lymphoid tissue (36, 37), and, in HIV-infected individuals, lower CD83 expression was observed in the spleen (38). Such observations may be explained by the impaired normal differentiation of DC in HIV infection described here. Recent studies of therapeutic vaccination in SIV-infected macaques using DC derived in vitro from autologous monocytes and pulsed with an inactivated virus preparation have shown promising humoral and T cell responses (39). These studies support previous work, showing that it is possible to generate functionally competent DC in vitro from monocytes of HIV-infected individuals (40, 41, 42). They also underline the failure in the ability of CD11c+ DC precursors in the blood to differentiate into effective APCs in HIV and SIV infection. Strategies designed to overcome the block in normal DC development may be worthy of investigation.

Our data reveal a novel differentiation pathway in the myeloid CD1c+ blood DC. An increased understanding of the properties of distinct mDC subsets may be important in the use of DC-based vaccines in the immunotherapy of cancer and infectious diseases including HIV-1 infection.

The authors have no financial conflict of interest.

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.

1

This work was supported by the Medical Research Council, United Kingdom.

3

Abbreviations used in this paper: DC, dendritic cell; mDC, myeloid DC; DC-SIGN, DC-specific ICAM-3-grabbing nonintegrin; SI, stimulation index.

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