Although CD34+ progenitor-derived immature dendritic cells (DCs) express CCR6, several recent studies reported that monocyte-derived immature DCs do not do so. We observed that DCs generated from monocytes in the presence of GM-CSF, IL-4, and TGF-β1 consistently responded to liver and activation-regulated chemokine (LARC, also known as macrophage inflammatory protein-3α). These immature DCs expressed one class of high-affinity binding sites for LARC, and expressed both CCR6 mRNA and protein. Therefore, LARC-CCR6 interaction presumably also contributes to the regulation of trafficking of monocyte-derived DCs, and utilization of TGF-β can potentially provide a ready source of CCR6+ monocyte-derived DCs for therapeutic purposes.

Dendritic cells (DCs)3 are professional APCs that play a critical role in the initiation of primary immune responses (1, 2). Upon the introduction of Ags into a host as by an infection, bone marrow-derived immature DCs migrate to the site of Ag deposition, take up and process Ags, and thereafter migrate, while undergoing a process of maturation, via the afferent lymphatic to regional draining lymph nodes to stimulate Ag-specific T lymphocytes (1, 2, 3, 4). The capacity of DCs to traffick through tissues to the lymph nodes is crucial to fulfill their roles in delivering Ags to T cells.

The signals controlling the traffic pattern of DCs are not fully elucidated, however, classic chemoattactants (fMLP, C5a, etc.) and chemokines are probably involved (2, 5, 6, 7, 8, 9). Chemokines, a superfamily of structurally related small proteins, contribute to regulating traffick of leukocytes (10, 11). Chemokine receptors belong to a subfamily of G-protein coupled seven-transmembrane domain receptors, and at least six of these chemokine receptors (CXCR4, CCR1, and CCR4–7) and their corresponding ligands have been shown to potentially participate in the control of DC migration (5, 6, 8, 9, 12, 13, 14, 15, 16, 17). A few of these ligands, in particular liver and activation-regulated chemokine (LARC) and EBI1 ligand chemokine (ELC, also known as macrophage inflammatory protein-3β), may be particularly important in directing DC trafficking because immature DCs express CCR6, the receptor for LARC, while mature DCs express CCR7, the receptor for ELC (14, 16, 17). LARC is largely detected within inflamed epithelial crypts in the periphery of tonsils, while ELC is found specifically in T cell-rich areas of tonsils (15). Therefore, the interaction of LARC with CCR6 is potentially involved in the recruitment of immature DCs to the site of Ag deposition, while the interaction of ELC with CCR7 is involved in attracting mature DCs to T cell-rich areas of secondary lymphoid tissues (15, 16, 17).

Human DCs can be generated in vitro both from CD34+ progenitors (18) and from CD14+ peripheral blood monocytes (Mo) (19, 20, 21). Although several laboratories have reported that Mo-derived DCs, unlike CD34+ progenitor-derived DCs, do not respond to LARC because they do not express the receptor (CCR6) for LARC (13, 14), we consistently observed that immature DCs generated from Mo in the presence of TGF-β1 responded to LARC. Furthermore, such immature Mo-derived DCs indeed express functional CCR6.

Recombinant human (rh) SDF-1α, GM-CSF (sp. act. = 107 U/mg), IL-4 (sp. act. = 2 × 106 U/mg), TGF-β1, and LARC were obtained from theNational Institutes of Health cytokine repository. Anti-CCR6 and anti-CD83 were purchased from R&D Systems (Minneapolis, MN) and Coulter-Immunotech (Miami, FL), respectively. The other Abs used were purchased from PharMingen (San Diego, CA).

Human peripheral blood Mo were isolated by Percoll gradient centrifugation as described previously (22). Mo were further purified (∼98%) by plastic adherence. Human T cells were purified from PBMCs by the use of human CD3 enrichment column (R&D Systems).

Mo were incubated in RPMI 1640 containing 10% FBS, 2 mM glutamine, 25 mM HEPES, 100 U/ml penicillin, and 100 μg/ml streptomycin, rhGM-CSF (50 ng/ml), rhIL-4 (50 ng/ml), and TGF-β1(5 ng/ml) at 37°C in humidified air with 5% CO2. The cultures were fed with the same cytokine-containing medium every two to three days. At day 7, rhTNF-α was added to a final concentration of 100 ng/ml to the cultures. The nonadherent cells harvested at days 7 and 9 were designated DC7 and DC9, respectively.

The migration of Mo and DCs was assessed using a 48-well microchemotaxis chamber technique as previously described (6). The results are presented as number of cells per high power field (No./HPF) or chemotactic index (C.I.). The statistical significance of the increase in cell migration was determined by unpaired t test.

DCs were first washed three times with FACS buffer (PBS, 1% FBS, 0.02% NaN3, pH 7.4), and then stained with various Abs at room temperature for 1 h. Subsequently, the cells were stained with FITC-conjugated goat anti-mouse IgG (Sigma, St. Louis, MO) for 30 min at room temperature, fixed with 1% paraformaldehyde in PBS, and analyzed with a flow cytometer (Coulter Epics).

Allogeneic MLR was performed as described (6) with minor modifications. Briefly, purified allogeneic T cells were cultured with Mo, DC7, or DC9 in a 96-well flat-bottom plate for 7 days at 37°C in humidified air with 5% CO2. The proliferative response was examined by pulsing the culture with [3H]TdR (0.5 μCi/well; NEN, Boston, MA) for another 18 h before harvesting. [3H]TdR incorporation was measured by a microbeta counter.

Equilibrium binding was performed in triplicate by adding constant amount of 125I-LARC (specific radioactivity = 2000 Ci/mmol; Amersham, Arlington Heights, IL) and increasing amounts of unlabeled LARC to individual 1.5-ml microfuge tubes, each containing 2 × 106 Mo or DCs suspended in RPMI 1640 containing 1% BSA, 2.5 mM HEPES, 0.05% NaN3. After incubation at 20°C with constant mixing for 1 h, the mixture was centrifuged through a 10% sucrose/PBS cushion, and the cell-associated radioactivity was measured with a 1227 Wallac gamma counter (Wallac, Gaithersburg, MD). Scatchard plotting was conducted by the use of Ligand.

Total RNA from Mo or DCs was isolated by the use of TRIzol Reagent (Life Technologies, Grand Island, NY). The RNAs were cleaned by RNase-free DNase I (Stratagene, La Jolla, CA) treatment. RT-PCR was performed by the use of GeneAmp RNA PCR Kit (Roche Molecular Systems, Branchburg, NJ). Forty and 26 cycles were used for the amplification of CCR6 and GAPDH sequences, respectively. The primers for CCR6 were 5′-CCCAAGCTTGGGGCGGGGAATCAATGAATTTCAGCGA-3′ and 5′-CCGCTCGAGCGGCTATCACATAGTGAAGGAGGACGCA-3′. The primers for human GAPDH were 5′-GATGACATCAAGAAGGTGGTGAA-3′ and 5′-GTCTTACTCCTTGGAGGCCATGT-3′. PCR products were identified on 1–2% agarose gel after ethidium bromide staining and were photodocumented.

Pilot studies showed that coculture of human Mo for 5–7 days with low (10 ng/ml) rather than high (50 ng/ml) concentrations of GM-CSF and IL-4 at times yielded DCs that exhibited a chemotactic response to LARC (data not shown). This suggested that the inability of Mo-derived DCs to respond to LARC might be due in part to use of higher concentrations of GM-CSF and IL-4 in previous studies, which aborted the immature stage of Mo-derived DCs (13, 14). To enrich the proportion of immature Mo-derived DCs, we added TGF-β1, a cytokine reported to arrest the maturation of CD34+ progenitor-derived DCs (23). DC7, generated from human Mo in the presence of 50 ng/ml of GM-CSF, IL-4, and 5 ng/ml of TGF-β1, consistently migrated in response to LARC, yielding a typical bell-shaped dose-dependent response with an optimal concentration at 100 ng/ml (Fig. 1,A). TGF-β1 by itself failed to induce DC generation (data not shown). Checkerboard analyses revealed that LARC-induced migration of DC7 was not due to chemokinesis (data not shown). Additionally, the chemotactic effect of LARC on DC7 was inhibited by pertussis toxin (Fig. 1 B), suggesting that the effect of LARC was mediated by a Gi-protein coupled receptor.

FIGURE 1.

Chemotaxis by LARC of DC7 cells generated from adherent Mo in the presence of TGF-β1. A, Dose-response result representative of >10 separate experiments is shown. B, DC7 cells were treated with (▦) or without (□) 100 ng/ml of pertussis toxin at 37°C for 30 min before adding to the chemotaxis chamber. All results are shown as mean ± SEM of triplicate wells.

FIGURE 1.

Chemotaxis by LARC of DC7 cells generated from adherent Mo in the presence of TGF-β1. A, Dose-response result representative of >10 separate experiments is shown. B, DC7 cells were treated with (▦) or without (□) 100 ng/ml of pertussis toxin at 37°C for 30 min before adding to the chemotaxis chamber. All results are shown as mean ± SEM of triplicate wells.

Close modal

To ensure that DCs differentiated from Mo in the presence of TGF-β1 were homogenous, we performed phenotypic analyses. Coculture of adherent Mo with GM-CSF, IL-4, and TGF-β1 for 7 days gave rise to nonadherent cells that are irregular in shape with a number of projecting dendrites typical of DCs (data not shown). Analyses of surface marker expression showed that these DC7 expressed little CD83, a moderate level of CD86, and a high level of HLA-DR (Fig. 2,A, upper panel). Furthermore, DC7 contained <1% of CD3+, CD14+, CD16+, CD19+, or CD62L+ cells (data not shown), suggesting that DC7 had little or no contamination with T cells, Mo, NK cells, B cells, or granulocytes. In contrast, DC7 contained 40% CD1a+, 92% CD11a+, and >99% CD11b+, CD13+, CD40+, and CD45+ cells (data not shown). The surface marker expression pattern of DC7 is quite similar to Mo-derived immature DCs generated in the absence of TGF-β1 (19, 20, 21, 24). Moreover, DC7 were inefficient in the stimulation of purified T cell proliferation in an allogeneic MLR (Fig. 2,B), further supporting the view that DC7 cells are immature DCs (19, 20, 24). On the other hand, DC9 expressed greater elevation of CD83, high levels of CD86 and HLA-DR (Fig. 2,A, lower panel), and acquired the capacity to stimulate the proliferation of pure allogeneic T cells (Fig. 2 B), characteristics of mature DCs (19, 20, 21, 24). The observation that immature DCs derived from Mo could be driven to mature by TNF-α even in the presence of TGF-β1 is also seen with CD34+ progenitor-derived DCs (23). Collectively, these results indicate that our DC7 and DC9 cells correspond to immature and mature Mo-derived DCs, respectively.

FIGURE 2.

Phenotypic and functional analyses of DC7 and DC9 cells. A, Expression of surface CD83, CD86, and HLA-DR by DC7 and DC9. Dotted line, cells stained with isotyped-matched control Abs. Solid line, cells stained with specific Abs as indicated. B, Allogeneic MLR. Mo, DC7, and DC9 were used at 5000/well while pure T cells were used at 105/well. C, Migration of Mo, DC7, and DC9 in response to LARC and SDF-1α. Cell migration (chemotactic index, C.I.) is expressed as mean ± SEM of each group. C.M. = chemotactic medium. ∗, p < 0.001 and ∗∗, p < 0.0001 when compared with C.M.

FIGURE 2.

Phenotypic and functional analyses of DC7 and DC9 cells. A, Expression of surface CD83, CD86, and HLA-DR by DC7 and DC9. Dotted line, cells stained with isotyped-matched control Abs. Solid line, cells stained with specific Abs as indicated. B, Allogeneic MLR. Mo, DC7, and DC9 were used at 5000/well while pure T cells were used at 105/well. C, Migration of Mo, DC7, and DC9 in response to LARC and SDF-1α. Cell migration (chemotactic index, C.I.) is expressed as mean ± SEM of each group. C.M. = chemotactic medium. ∗, p < 0.001 and ∗∗, p < 0.0001 when compared with C.M.

Close modal

CD34+ progenitor-derived immature DCs respond, while mature DCs lose their responsiveness, to LARC (15). Fig. 2,C similarly shows that only DC7 (immature), but not Mo or DC9 (mature), migrated toward LARC. The inability of Mo and DC9 to migrate toward LARC was not due to reduced motility of the cells because they migrated toward 100 ng/ml of SDF-1α equally as well as DC7 cells (Fig. 2 C). The difference in chemotactic response of Mo, DC7, and DC9 cells toward LARC occurred despite the fact that these cells were derived from the same batch of adherent monocytes. Consequently, immature Mo-derived DCs can respond to LARC at a similar developmental stage as do CD34+ progenitor-derived DCs (15).

To identify the receptor(s) that mediate the effect of LARC on DC7, we conducted equilibrium binding experiments. The results show that only DC7, but neither Mo nor DC9, specifically bound 125I-LARC (Fig. 3). Furthermore, the binding of 125I-LARC to DC7 was competitively inhibited by addition of increasing amount of unlabeled LARC. Scatchard plot analysis demonstrated that DC7 cells possess only one class of specific LARC-binding sites (about 42,000 sites/cell) with high affinity. The Kd of LARC-DC7 binding is 1.6 nM, which lies in the range of previously reported Kd values for binding of LARC to CCR6 (13, 14, 25).

FIGURE 3.

Inhibition by LARC of 125I-LARC binding to Mo, DC7, and DC9 cells. Approximately 6000 cpm of 125I-LARC was added into each tube of cell suspension. Inset, Scatchard plot analysis of the inhibition binding data of DC7. Kd, equilibrium dissociation constant. R, LARC-binding sites/DC7. Two separate experiments gave rise to similar results.

FIGURE 3.

Inhibition by LARC of 125I-LARC binding to Mo, DC7, and DC9 cells. Approximately 6000 cpm of 125I-LARC was added into each tube of cell suspension. Inset, Scatchard plot analysis of the inhibition binding data of DC7. Kd, equilibrium dissociation constant. R, LARC-binding sites/DC7. Two separate experiments gave rise to similar results.

Close modal

The possibility that Mo-derived DC7 generated in the presence of TGF-β1 express CCR6 was investigated by RT-PCR. Indeed, DC7 expressed a significant level of CCR6 mRNA, but this was not the case for Mo or DC9 derived from the same donor (Fig. 4,A, upper panel). The amplified CCR6 fragment was displayed on agarose gel as a band of 1150 bp, which was very close to the anticipated size (1149 bp). Amplification of GAPDH mRNA by the use of identical volumes of reverse-transcribed RNA samples as used in the amplification of CCR6 gave rise to similar GAPDH cDNA bands for Mo, DC7, and DC9 (Fig. 4,A, middle panel), indicating that identical amounts of RNAs were used. To confirm the specificity of PCR for the amplification of CCR6 mRNA, the cDNA bands depicted in the upper panel of Fig. 4,A were purified, digested with EcoRI, and fractionated on agarose gel (Fig. 4,A, lower panel). The generation of two fragments identical with the anticipated sizes confirmed that the band displayed in the upper panel was indeed CCR6 cDNA. When DC7 and DC9 cells were stained with anti-CCR6 Abs and analyzed by FACScan, DC7, but not DC9, cells showed a significant shift with an ∼10-fold increase in mean fluorescence intensity (Fig. 4 B), demonstrating clearly that Mo-derived immature DCs also express CCR6 at the protein level.

FIGURE 4.

Expression by DC7 cells of CCR6 at both mRNA and protein levels. A, RT-PCR products of CCR6 (upper panel) and GAPDH (middle panel) displayed by 1% agarose gel electrophoresis. The anticipated size for CCR6 and GAPDH is 1149 bp and 246 bp, respectively. The lower panel displays the production of two fragments with anticipated sizes from EcoRI digestion of CCR6 cDNA bands depicted in the upper panel. Marker, 1 kb DNA ladder. B, Overlay histograms of DC7 and DC9 cells stained with either isotype-matched control (dotted line) or anti-CCR6 Abs.

FIGURE 4.

Expression by DC7 cells of CCR6 at both mRNA and protein levels. A, RT-PCR products of CCR6 (upper panel) and GAPDH (middle panel) displayed by 1% agarose gel electrophoresis. The anticipated size for CCR6 and GAPDH is 1149 bp and 246 bp, respectively. The lower panel displays the production of two fragments with anticipated sizes from EcoRI digestion of CCR6 cDNA bands depicted in the upper panel. Marker, 1 kb DNA ladder. B, Overlay histograms of DC7 and DC9 cells stained with either isotype-matched control (dotted line) or anti-CCR6 Abs.

Close modal

Collectively, our data demonstrate that DCs generated in the presence of TGF-β1 are immature and express a single class of sites for LARC as well as its receptor, CCR6. The discrepancies between our data and those reported by others (13, 14) can be attributed to differences in culture conditions, particularly the addition of TGF-β1. Mo-derived DCs generated in the presence of TGF-β1 are phenotypically indistinguishable from those generated in the absence of TGF-β1 (19, 20, 21, 24). However, they are “truly” immature in terms of their total inability to stimulate an allogeneic MLR (Fig. 2 B) in comparison with Mo-derived DCs generated in the presence of only GM-CSF and IL-4 that exhibit some, albeit low, capacity to stimulate the proliferation of allogeneic T cells (20, 21, 24). Thus, DC7 generated in the absence of TGF-β1 may already be undergoing maturation and therefore no longer express CCR6.

Considering that the usage of ex vivo Ag-loaded autologous DCs for vaccination is one of the most promising approaches to initiate antiviral and antitumor immunity, our findings may prove valuable in providing a means of generating Mo-derived immature DCs using TGF-β1. Therefore, the optimal means of generating Ag-loaded Mo-derived DCs in vitro for in vivo vaccination might be to generate “truly” immature DCs using GM-CSF, IL-4, and TGF-β1 to pulse them with target Ags and subsequently to induce them to fully mature before in vivo administration.

We thank Drs. E. L. Nelson (Clinic Services Program, Science Applications International Corp, Frederick, MD) and F. Ruscetti (National Cancer Institute-Frederick Cancer Research and Development Center, Frederick, MD) for providing us with PE-conjugated anti-CD83 and human TGF-β1, respectively. We also thank N. Dunlop for technical assistance. The secretarial assistance of Ms. C. Fogle is gratefully appreciated.

1

The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government. The publisher or recipient acknowledges right of the U.S. Government to retain a nonexclusive, royalty-free license in and to any copyright covering the article. D.Y. is supported in part by a fellowship from the Office of International Affairs, National Cancer Institute, National Institutes of Health. O.M.Z.H. is funded in part by the National Cancer Institute, National Institutes of Health Contract N01-CO-56000.

3

Abbreviations used in this paper: DC, dendritic cells; Mo, monocyte; rh, recombinant human; LARC, liver and activation-regulated chemokine; ELC, EBI1 ligand chemokine; SDF-1α, stromal-derived factor 1α.

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