Differential Regulation of Vitamin D Receptor and Its Ligand in Human Monocyte-Derived Dendritic Cells

Generation of moDCs

PBMCs were derived from buffy coats of healthy donors by density centrifugation over Lymphoprep (Robbins Scientific, Sunnyvale, CA), resuspended in RPMI 1640/2% human AB serum, and allowed to adhere to plastic at 37°C for 2 h. Nonadherent cells were then removed and monocytes were cultured in IMDM (Life Technologies, Rockville, MD) plus 5% human AB serum (HD Supplies, Buckinghamshire, U.K.) plus 2 mM l-glutamine plus 100 μg/ml streptomycin plus 100 μg/ml penicillin for 7 days in the presence of 800 U/ml GM-CSF (Novartis Pharmaceuticals, East Hanover, NJ) and 1000 U/ml IL-4 (R&D Systems, Abingdon, U.K.) in 75-cm² flasks (Iwaki, Tokyo, Japan). In parallel experiments, moDCs were propagated under serum-free conditions in macrophage serum-free media (Life Technologies). 1α25(OH)₂D₃ (10⁻⁸ M), 25(OH)D₃ (10⁻⁸–10⁻⁷ M), or vehicle (Sigma-Aldrich, St. Louis, MO) were added in some experiments either at the beginning of culture or at time points indicated in the text. Media were replenished at 3-day intervals. Immature DC were resuspended in the above media ± vitamin D₃ metabolites (on day 7) and incubated in 12-well plates at a concentration of 5 × 10⁵/ml and matured for 48 h in the presence of one of the following maturation stimuli: 5 μg/ml G28.5 CD40 Ab (kindly provided by Prof. J. Gordon, Department of Immunology, University of Birmingham, Birmingham, U.K.) plus 5 μg/ml goat anti-mouse Ig Fc cross-linker (Pierce, Rockford, IL), 50 ng/ml TNF-α (Peprotech, London, U.K.), 1000 U/ml IFN-γ, 1 μg/ml LPS Escherichia coli, or 20 μg/ml poly I:C (all obtained from Sigma-Aldrich). In control experiments, 5 μg/ml nonspecific mouse IgG1 MOPC21 (Sigma-Aldrich) plus 5 μg/ml goat anti-mouse Ig Fc cross-linker (Sigma-Aldrich) or PBS were substituted for the various maturation stimuli. In experiments to test the regulation of 1αOHase, 20 μM p38 mitogen-activated protein kinase (MAPK) inhibitor SB203580 (Sigma-Aldrich) or 50 μM of the NF-κB inhibitor N-α-tosyl-l-phenylalanine chloromethyl ketone (TLCK; Sigma-Aldrich) were added 60 min before the maturation stimulus.

Flow cytometric analysis

The cell surface phenotype of moDCs or DCs propagated in the presence of vitamin D₃ metabolites was analyzed using the following mAbs: anti-CD1a-fluorescecin isothiocyanate (HI149;IgG1), anti-CD14-PE (M5E2. IgG2a), anti-CD83-PE (HB15E, IgG1), anti-CD86-PE (IT2.2, IgG2b), anti-HLA-DR-Cychrome (G46.6, IgG2a; San Diego, CA), and the isotype controls labeled with the appropriate fluorochrome (MOPC21, IgG1; G155–178, IgG2a; and 27–35, IgG2b) were obtained from BD PharMingen (San Diego, CA). The staining of the samples was measured on a FACS Coulter Epics XL (Beckman Coulter, Fullerton, CA), using System II software (Beckman Coulter), and was analyzed with WinMDI software (from Prof. J. Trotter, The Scripps Research Institute, San Diego, CA).

Cytokines

In experiments to test the effect of vitamin D₃ metabolites upon IL-12 production by moDCs, day 7 DCs were incubated for 48 h in 48-well plates at a concentration of 5 × 10⁵/ml with CD40 ligand (CD40L)-transfected or control nontransfected L cells at a ratio of 2:1. IL-12p70 and IL-12p40 levels were evaluated by ELISA kits according to the manufacturer’s instructions (BD PharMingen). The IL-12p40 kit detects the IL-12p40 homodimer or monomer but not the bioactive IL-12p70 heterodimer.

Cytochemistry staining

Cytospins were prepared from monocytes, immature moDC, and mature moDC, washed with PBS and stained for 1 h at room temperature with sheep anti-1αOHase Ab (12), which had been diluted 1/200. After washing, the slides were incubated with secondary biotinylated donkey anti-sheep Ab (Binding Site, Birmingham, U.K.) for 30 min, followed by a second 30-min incubation with a streptavidin-biotin complex (strepABComplex; DAKO, Cambridgeshire, U.K.). The staining was visualized using 3′-diaminobenzidine tetrahydrochloride (Sigma-Aldrich). Finally, the cells were counterstained with Mayer’s hematoxylin. The percentage of stained cells and intensity of staining were measured by counting a minimum of 200 cells per slide. All analyses were performed single blind.

EMSAs

These were performed as previously described (18). For the detection of NF-κB binding activity, an oligonucleotide containing the HIV-long terminal repeat NF-κB consensus binding sequence (5′-GAT CAG GGA CTT TCC GCT GGG GAC TTT CC-3′) was used.

Phospho-p38-immunoblots

Phospho-p38 immunoblots were performed as described previously (18) using a phosphospecific p38 MAPK (Thr¹⁸⁰/Tyr¹⁸²) Ab (New England Biolabs, Beverly, MA) and were compared with total p38 MAPK.

Quantitative RT-PCR analysis of 1α-OHase, VDR, and 25(OH)D₃ 24-hydroxylase (24(OH)ase) mRNA expression

1αOHase, VDR, and 24(OH)ase mRNA levels were analyzed using an ABI 7700 sequence detection system (PE Biosystems, Warring, U.K.) using previously reported protocols (12). RNA was extracted from cell pellets using the StrataPrep total RNA miniprep kit (Stratagene, Amsterdam, The Netherlands) and was reverse-transcribed using AMV reverse transcriptase (Promega, Madison, WI) and random hexamers in 40-μl reaction volumes according to manufacturer’s instructions. Amplification of cDNAs was performed in 25-μl volumes on 96-well plates, in a reaction buffer containing TaqMan Universal PCR Master Mix, 3 mM Mn(Oac)₂, 200 μM dNTPs, 1.25 U AmpliTaq Gold polymerase, 1.25 U AmpErase UNG, 150 nmol TaqMan probe, 900 nmol primers, and 50 ng of cDNA. All reactions were multiplexed with the housekeeping gene 18S rRNA, provided as a preoptimized control probe (PE Biosystems) enabling data to be expressed in relation to an internal reference to allow for differences in sampling and RT efficiency. Data were obtained as cycle threshold (Ct) values (the cycle number at which logarithmic PCR plots cross a calculated threshold line) according to the manufacturer’s guidelines and were used to determine ΔCt values (ΔCt = Ct of the target gene − Ct of the housekeeping gene). Measurements were conducted in triplicate for each individual experiment. All PCRs were conducted with the primers and probes outlined below using the following reaction conditions: 50°C for 2 min, 95°C for 10 min; 44 cycles of 95°C for 15 s; 60°C for 1 min.

Primers and probes for PCRs

PCR primer and probe sequences for 1αOHase were as follows: forward primer, 5′-CACCCGACACGGAGACCTT-3′; reverse primer, 5′-TCAACAGCGTGGACACAAACA-3′ and TaqMan probe, 5′-TCCGCGCTGTGGGCTCGG-3′. For 24(OH)ase: forward primer, 5′-CAAACCGTGGAAGGCCTATC-3′; reverse primer, 5′-AGTCTTCCCCTTCCAGGATCA-3′ and Taqman probe, 5′-ACTACCGCAAAGAAGGCTACGGGCT-3′. For VDR: forward primer, 5′-CTTCAGGCGAAGCATGAAGC-3′; reverse primer, 5′-CCTTCATCATGCCGATGTCC-3′ and TaqMan probe, 5′-AAGGCACTATTCACCTGCCCCTTCAA-3′. In each case, the housekeeping gene used to derive ΔCt values was 18S rRNA which was analyzed using primers and probes provided by the manufacturer (PE Biosystems).

1αOHase activity

Cells were washed in serum-free RPMI 1640 and were then resuspended at 1 × 10⁶ cells/ml. Aliquots of this suspension (400 μl) were then incubated with 10 nM [³H]25(OH)D₃ (specific activity 180 Ci/mmol; Amersham, London, U.K.) for 4 h at 37°C. The reaction was terminated by freezing at −20°C. Cell extracts and medium were combined and vitamin D metabolites were extracted in 2.5 ml of chloroform:methanol (4:1 vol:vol). Following evaporation of the organic phase, steroids were resuspended in 50 μl of dichloromethane:isopropanol (9:1 vol:vol). Standard lanes were included which contained only [³H]25(OH)D₃ or [³H]1α25(OH)₂D₃. Production of [³H]1α25(OH)₂D₃ was measured on a Bioscan System 200 imaging TLC plate scanner (Bioscan, Edmonds, WA). Data were reported as fmol of 1α25(OH)₂D₃ produced per hour per 10⁶ cells.

Analysis of nuclear binding of 1α25(OH)₂D₃

Numbers of intracellular receptors for 1α25(OH)₂D₃ were assessed by whole cell nuclear association assays. Briefly, cells were resuspended in serum-free RPMI 1640 to give 0.5 × 10⁷ cells/ml. Aliquots (100 μl) of these suspensions were then added to glass tubes containing a single saturating dose (5 nM) of [³H]1α25(OH)₂D₃ (specific activity 180 Ci/mmol; Amersham) in the presence or absence of a 200-fold excess of unlabeled 1α25(OH)₂D₃ (to determine nonspecific binding). Cells were then incubated for 1 h to allow association of ligand-VDR complexes with nuclei and then washed twice with 1 ml of PBS at 4°C to remove unincorporated [³H]1α25(OH)₂D₃. The final cell pellets were incubated with 0.5 ml of lysis buffer (0.25 M sucrose, 20 mM HCl, 1.1 mM magnesium chloride, and 0.5% Triton X-100, pH 7.4) and centrifuged at 1000 × g for 5 min. This step was repeated and the resulting crude nuclear pellet was resuspended in 100 μl of PBS and 100 μl of ethanol, and transferred to 4 ml of scintillation fluid before scintillation counting. Maximal binding was determined as fmol 1α25(OH)₂D₃ bound per 10⁶ cells and then converted to VDR per cell using Avogadro’s constant.

Data analysis

Statistical analysis was performed using one-way or repeated measure ANOVA, with posthoc testing using the Student Newmans Keuls multiple comparison posttest (Instat version 2.04a; GraphPad Software, San Diego, CA).

Human moDCs synthesize 1α25(OH)₂D₃

To determine whether human DCs are able to synthesize 1α25(OH)₂D₃, monocytes were incubated in the presence of IL-4 and GM-CSF for 7 days and were then matured by CD40 ligation for 48 h. At various time points, RNA was prepared and real time RT-PCR was performed to determine 1αOHase mRNA expression. Differentiation and maturation of moDCs was accompanied by a marked increase in 1αOHase mRNA expression as compared to the starting population, increasing >10-fold in day 7 immature moDCs and >60-fold in moDC matured by CD40 ligation (Fig. 1⇓A). The amplified product corresponded to the expected size and sequence indicating that, in common with macrophages, the transcript is identical to that detected in proximal renal tubular cells (data not shown). We also examined the effect of other maturation stimuli representative of both T cell and microbial origin for their ability to induce the transcriptional up-regulation of 1αOHase. As shown in Fig. 1⇓B, under the conditions tested, CD40 ligation, TNF-α, IFN-γ, poly I:C, and LPS treatments of day 7 immature moDCs were associated with ∼5- to 12-fold induction of 1αOHase as compared with nontreated controls. Immunocytochemical staining of monocytes, immature and mature moDCs for the 1αOHase enzyme demonstrated a similar increase in protein expression, with dense staining, in particular of mature moDCs (Figs. 1⇓C and 3B). In parallel, 1α25(OH)₂D₃ synthesis, as estimated by thin layer chromatography of cellular extracts incubated in the presence of the [³H]25(OH)D₃ precursor, was absent in monocytes and increased dramatically following differentiation into immature moDC (Fig. 1⇓, D and E). CD40 ligation-induced moDC maturation further boosted 1α25(OH)₂D₃ synthesis by >3-fold as compared with immature moDCs or mock-treated cells (Fig. 1⇓, D and E, and data not shown). moDC 1αOHase mRNA expression and 1α25(OH)₂D₃ synthesis was similar to that of GM-CSF-propagated macrophages before and following activation (Fig. 1⇓E and data not shown). In additional experiments, we confirmed that moDC generated under serum-free conditions also up-regulated 1αOHase gene transcription and activity, indicating that this phenomenon is reproducible under alternate culture conditions (data not shown). Thus, moDCs develop the capacity to synthesize 1α25(OH)₂D₃ in vitro to an extent which is comparable to macrophages and which is enhanced upon terminal differentiation induced by a diverse set of stimuli.

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FIGURE 1.

Human moDCs synthesize 1α25(OH)₂D₃. A, Monocytes were cultured in 1000 U/ml IL-4 and 800 U/ml GM-CSF for 7 days and then immature moDCs were matured by CD40 ligation for 48 h. RNA was extracted from cells at the indicated time points and real time RT-PCR was performed to evaluate 1αOHase expression. Mean fold change (±SEM) in 1αOHase mRNA expression (y-axis) compared with monocytes vs day of culture (x-axis) is shown. Data shown are for eight independent experiments. Significant differences were evident between day 1 and both days 7 and 9 (p < 0.001 both comparisons) and between days 7 and 9 (p < 0.01). B, moDCs were propagated as above and on day 7 were left untreated or matured with CD40 mAb, 50 ng/ml TNF-α, 1000 U/ml IFN-γ, 1 μg/ml LPS E. coli, or 20 μg/ml poly I:C for 48 h. Maturation with each stimulus was confirmed by flow cytometric staining for CD83 (not shown). Mean fold increases (±SEM) in 1αOHase mRNA expression compared with nontreated cells are shown. Significant increases were observed between nontreated cells and all maturation stimuli tested (CD40 mAb, TNF-α, LPS (p < 0.001), IFN-γ, poly I:C (p < 0.01)). C, Photomicrographs (×400) of immunocytochemical staining of cytospins prepared from monocytes, day 7 immature moDCs, and TNF-α-matured moDC using polyclonal antisera directed against 1αOHase. Negative staining of cytospins incubated without primary Ab or with an excess of the peptide to which the primary was directed confirmed the specificity of staining (not shown). Data are representative of four independent experiments performed. D, Thin layer chromatographic analysis of products following incubation of 1 × 10⁶/ml CD40 ligation-matured moDCs with 10⁻⁸ M [³H]25(OH)D₃ for 4 h at 37°C. Graph shows disintegrations per minute (y-axis, dpm) vs migration (x-axis, mm). Positions of 1α25(OH)₂D₃ (a) and 25(OH)D₃ (b) as evaluated by running standards in parallel assays are shown. Data shown are representative of three independent experiments performed. E, Monocytes were cultured in 1000 U/ml IL-4 and 800 U/ml GM-CSF or 800 U/ml GM-CSF alone for 7 days to generate immature DCs or macrophages, respectively, and then were activated by CD40 ligation. The percentages of cell staining for CD14 were 3.3 and 69.5% on day 7 immature moDCs and macrophages, respectively. Evaluation of 1αOHase was performed as above and the mean (±SEM triplicate values) of 1αOHase activity in fmol 1α25(OH)₂D₃ synthesis per 10⁶ cells per hour is shown. The data shown is representative of three independent experiments.

25(OH)D₃ inhibits DC differentiation, maturation, and allostimulatory potential

Loss of the 1α-hydroxyl group from 1α25(OH)₂D₃ is associated with an ∼500-fold reduction in the binding affinity of ligand to VDR (19) and, thus, no VDR-mediated biological activity is demonstrable at physiological concentrations of 25(OH)D₃ (20). The ability of human moDCs to generate 1α25(OH)₂D₃ from the inactive precursor 25(OH)D₃ suggested that the latter could have inhibitory effects upon the differentiation or function of human moDCs. To test this hypothesis we exposed moDC cultures from the first day of culture to 10⁻⁸–10⁻⁷ M (10–100 nM) 25(OH)D₃, representative of concentrations below or within the normal physiological range (19) and evaluated the effect upon DC differentiation, maturation, allostimulatory potential, and survival. In the same experiments, as a positive control, DCs were propagated in the presence of 10⁻⁸ M (10 nM) 1α25(OH)₂D₃, the concentration at which maximal inhibitory effects are observed (5, 6). By day 7, immature DCs cultured in the presence of GM-CSF and IL-4 were CD14-negative and had acquired surface CD1a expression (Fig. 2⇓, A and B). As in previous studies, DCs differentiated in the presence of 10⁻⁸ M 1α25(OH)₂D₃ showed a marked decrease in CD1a expression, while CD14 remained elevated. Addition of 10⁻⁸–10⁻⁷ M 25(OH)D₃ at the initiation of culture also resulted in marked inhibition of CD1a expression at all concentrations tested. Furthermore, we observed a concentration-dependent failure to down-regulate CD14 with relatively minor effects observed at 10⁻⁸ 25(OH)D₃ but significant inhibition at 10⁻⁷ M 25(OH)D₃, equivalent to that observed in the presence of 10⁻⁸ M 1α25(OH)₂D₃. Consistent with this, day 7 immature moDCs that had been cultured in the presence of 10⁻⁸–10⁻⁷ M 25(OH)D₃ had impaired allostimulatory function as compared with controls (data not shown). As shown in Fig. 2⇓, C and D, and in accordance with other studies (5, 6), DCs generated in the presence of 10⁻⁸ M 1α25(OH)₂D₃ failed to up-regulate CD83 expression in response to CD40 ligation to the same degree as controls. Again, similar concentration-dependent effects were observed following the addition of 10⁻⁸–10⁻⁷ M 25(OH)D₃, with greater inhibition at 10⁻⁷ M 25(OH)D₃, equivalent to that observed in the presence of10⁻⁸ M 1α25(OH)₂D₃. In addition, DCs cultured in the presence of 10⁻⁸ M 1α25(OH)₂D₃ and higher concentrations of 25(OH)D₃ both failed to up-regulate other molecules involved in the development of accessory functions, including CD80, CD86, and HLA-DR (Fig. 2⇓, C and D). Similar results were observed when DCs were matured in the presence of TNF-α (data not shown). Furthermore, as compared to controls, DCs differentiated in the presence of both 10⁻⁸ M 1α25(OH)₂D₃ and 10⁻⁷ M 25(OH)D₃ demonstrated an almost complete failure in their capacity to produce IL-12p70 and IL-12p40 in response to CD40 ligation (Fig. 2⇓E) and accordingly had weak APC activity in an allogeneic mixed leukocyte reaction (Fig. 2⇓F). At the concentrations used and under the specific conditions described above, we observed no effects of 25(OH)D₃ upon day 9 DC viability as evaluated by annexin V-FITC (FITC)-propidium iodide (PI) staining (control-FITC⁻PI⁻ 82.4 ± 3.4%; 10⁻⁸ M 1α25(OH)₂D₃-FITC⁻PI⁻ 80.0 ± 3.0%; and 10⁻⁷ M 25(OH)D₃- FITC⁻PI⁻ 84.0 ± 1.6%, n = 6). In conclusion, these results suggest that the active synthesis of 1α25(OH)₂D₃ from its inactive precursor restricts the differentiation of immature moDCs in vitro and their subsequent sensitivity to maturation stimuli. This phenomenon appears to operate at physiological concentrations of 25(OH)D₃.

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FIGURE 2.

25(OH)D₃ inhibits DC differentiation, maturation, and allostimulatory potential. A, Monocytes were cultured in 1000 U/ml IL-4 and 800 U/ml GM-CSF for 7 days. 1α25(OH)₂D₃ (10⁻⁸ M, 10 nM), 25(OH)D₃ (10⁻⁸–10⁻⁷ M, 10–100 nM), or vehicle was added at the start of culture. On day 7, the percentage of cells staining positively for cell surface CD1a (top graph) and CD14 (bottom graph) was evaluated. Results shown are representative of six independent experiments. For CD1a, significant differences were evident between controls and 1α25(OH)₂D₃ (p < 0.001) and all concentrations of 25(OH)D₃ tested (p < 0.001). For CD14, significant differences were evident between controls and 1α25(OH)₂D₃ (p < 0.05) and at 10⁻⁷ M 25(OH)D₃ (p < 0.05). B, Representative flow cytometric dot plots for cell surface CD1a and CD14 staining of day 7 DCs cultured in the presence of vehicle, 1α25(OH)₂D₃ (10⁻⁸ M) or 25(OH)D₃ (10⁻⁷ M). Percentage of positive cells, set according to isotype controls, are shown in each quadrant. C, Histogram to show inhibitory effects of differing concentrations of 25(OH)D₃ (10⁻⁸–10⁻⁷ M, 10–100 nM) as compared with 1α25(OH)₂D₃ (10⁻⁸ M, 10 nM) upon membrane phenotype of day 9 moDCs/vitamin D₃ metabolite-treated moDCs matured by CD40 ligation. Percentage inhibition of mean cellular fluorescence (MCF) = (1 − (MCF test/MCF control)) × 100. D, Flow cytometric histograms showing staining for CD83, CD80, CD86, and HLA-DR for day 9 moDCs/vitamin D₃ metabolite-treated moDCs matured by CD40 ligation (isotype control, open histograms; day 9 DC, filled histograms). Figures in the upper right corner of each histogram represent MCF of day 9 DCs stained with specific fluorochrome-labeled Ab minus that of fluorochrome-labeled isotype controls. Results shown are representative of at least six independent experiments, except for CD80 which was evaluated in three independent experiments with similar results. E, moDCs were cultured in the presence of vehicle, 1α25(OH)₂D₃ (10⁻⁸ M), or 25(OH)D₃ (10⁻⁷ M) and on day 7 were cultured in the presence of CD40L-transfected cells or nontransfected control cells. At 48 h, culture supernatants were removed and IL-12p70 and IL-12p40 concentrations were measured using an ELISA method. Histogram shows mean ± SEM IL-12p70 (▪, n = 4) or IL-12p40 (□, n = 3) for each condition. Concentration of IL-12 (picograms per milliliter) = IL-12 concentration in the presence of CD40L transfectants − IL-12 concentration in the presence of nontransfected controls. Significant differences were evident between control DCs and 1α25(OH)₂D₃ (IL-12p70, p < 0.001; IL-12p40, p < 0.05) and 25(OH)D₃ (IL-12p70, p < 0.001; IL-12p40, p < 0.05). F, Day 9 moDCs/vitamin D₃ metabolite-treated moDCs matured by CD40 ligation were extensively washed, irradiated 3000 rad, and 1.5 × 10⁴ cells and doubling dilution thereof were added to 96-well round bottom plates. Purified allogeneic 10⁵ CD4⁺CD45RA⁺ T cells were added to each well and the cells were incubated for 5 days. Cell proliferation was assessed by [³H]thymidine incorporation after cells were pulsed with [³H]thymidine for the last 16 h of culture. All assays were performed in triplicate. Graph shows proliferation (cpm, y-axis) vs number of moDCs per well (x-axis). Result is representative of six independent experiments. Similar results were obtained when moDCs or vitamin D₃ metabolite-treated moDCs were matured in the presence of TNF-α.

Regulation of 1αOHase

The above data are indicative of a link between immature moDC maturation and the up-regulation of 1αOHase expression. Integral to the maturation program of DCs in response to divergent inflammatory stimuli is the activation and cooperative interaction of the p38 MAPK and NF-κB pathways (21, 22). Therefore, we reasoned that specific inhibition of each of these pathways would also inhibit the up-regulation of 1αOHase expression. As shown in Fig. 3⇓, A and B, inhibition of the p38 MAPK pathway using the pyridinyl imidazole compound SB203580 resulted in a major reduction in the expression of the 1αOHase induced by TNF-α as demonstrated by immunocytochemical staining. Western blots of total cell lysates were performed in parallel to confirm that SB203580 inhibited p38 MAPK phosphorylation (Fig. 3⇓C). Furthermore, inhibition of nuclear NF-κB translocation using the proteosomal inhibitor TLCK was also associated with a failure to up-regulate 1αOHase expression induced by TNF-α (Fig. 3⇓, A and B). We confirmed in these experiments that TLCK effectively inhibited TNF-α-induced increases in nuclear NF-κB binding (Fig. 3⇓D). In contrast, SB203580 had no effect upon nuclear NF-κB binding, confirming that under the conditions used in these experiments, its effects were limited to inhibition of the p38 MAPK pathway alone. We confirmed, as reported previously (23), that inhibition of both these pathways inhibited up-regulation of cell surface CD83, but had no effect on moDC viability (data not shown). In additional experiments (Fig. 3⇓E), we confirmed that disabling either the p38 MAPK or NF-κB pathways led to almost complete inhibition of the transcriptional up-regulation of 1αOHase upon maturation.

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FIGURE 3.

Regulation of 1α(OH)ase. A, Photomicrographs (×400) of immunocytochemical staining of cytospins prepared from TNF-α-matured moDC cultured in the presence or absence of TLCK (50 μM) or SB203580 (20 μM) using polyclonal antisera directed against 1αOHase. Data are representative of three independent experiments performed. B, Stained cytospins were scored for the number of densely staining cells (defined as cytoplasmic staining that obscures the nuclear outline). All analyses were performed single blind and a minimum of 200 cells were counted. Mean percentage (±SEM) of densely staining monocytes, day 7 immature moDCs, and day 9 moDCs matured with TNF-α (50 ng/ml) in the presence or absence of TLCK or SB203580 are shown for three independent experiments. C, Immature moDCs were treated for 60 min with SB203580 (20 μM) or were mock-treated before the addition of 50 ng/ml TNF-α. After 30 min, cell lysates were prepared and subjected to immunoblotting using mAbs specific for total (p38) or phospho-p38 MAPK (p-p38) as indicated by the arrows. D, Immature moDCs were treated for 60 min with TLCK (50 μM) or SB203580 (20 μM) or mock-treated, before the addition of TNF-α (50 ng/ml). After 30 min, nuclear extracts were prepared and nuclear NF-κB activity was determined (arrow). E, Immature moDCs were left untreated, or were matured by CD40 ligation with or without the prior addition 60 min before of TLCK (50 μM), SB203580 (20 μM), or 1α25(OH)₂D₃ (10⁻⁸ M). RNA was extracted and real time RT-PCR was performed to evaluate 1αOHase expression. Mean fold change (±SEM) in 1αOHase mRNA expression compared with untreated cells is shown; three independent experiments were performed. F, moDCs were prepared as above, with or without the addition separately of 1α25(OH)₂D₃ (10⁻⁸ M) or 25(OH)D₃ (10⁻⁷ M). RNA was extracted at the indicated time points and real time RT-PCR was performed to evaluate 24(OH)ase mRNA expression. Similar results were obtained in a second experiment performed.

In proximal renal tubule cells, 1αOHase gene expression is subject to exquisite feedback inhibition upon exposure to 1α25(OH)₂D₃, an effect that is dependent upon VDR expression (24). To test the effect of 1α25(OH)₂D₃ upon 1αOHase gene expression, immature moDCs were matured by CD40 ligation in the presence or absence of 10⁻⁸ M 1α25(OH)₂D₃. As shown in Fig. 3⇑E, 1α25(OH)₂D₃ had little effect upon the expression of 1αOHase mRNA. The lack of effect was also observed if 1α25(OH)₂D₃ was added >24 h before the application of the maturation stimulus, indicating that this was not due to slower kinetics of inhibition (data not shown). The resistance of 1αOHase gene expression to the negative regulatory effects of 1α25(OH)₂D₃ is also observed in other cells derived from extrarenal sites, such as keratinocytes (25) or macrophages (26). In these examples, the synthesis of 1α25(OH)₂D₃ may be controlled by the competing effects of another enzyme, 24(OH)ase, which inactivates 1α25(OH)₂D₃. 24(OH)ase is itself induced by 1α25(OH)₂D₃ through a VDR-mediated mechanism (27, 28). As shown in Fig. 3⇑F, in the absence of 1α25(OH)2D₃, human moDCs do not express significant levels of 24(OH)ase. In contrast, exposure of differentiating moDCs to 1α25(OH)₂D₃ increased 24(OH)ase gene expression dramatically (up to 3000-fold) at early time points (day 3 of culture), but notably the degree of induction then fell by ∼6-fold on days 7 and 9. This fall in 24(OH)ase expression did not represent the consequence of 24(OH)ase-mediated metabolism of 1α25(OH)2D₃, because the active ligand was continually replenished in the medium. Importantly, 25(OH)D₃ had little effect upon 24(OH)ase gene expression on day 3, but consistent with its conversion to active ligand, induced 24(OH)ase expression at later time points to a degree similar to exogenous 1α25(OH)₂D₃.

In summary, 1αOHase up-regulation in moDCs is associated with the p38 MAPK-and NF-κB-dependent maturation of DCs, but its transcription is resistant to the negative regulatory effects of 1α25(OH)₂D₃. A degree of control for 1α25(OH)₂D₃ synthesis may be imparted by the induction of an inactivating enzyme, 24(OH)ase, particularly at early time points during differentiation.

VDR expression is down-regulated during DC differentiation

A major arbiter of the dendritic cellular response to either exogenously or endogenously derived 1α25(OH)₂D₃ ligand will be the level of expression of VDR and/or the presence of coactivators that are recruited to the nuclear receptor complex. One indirect measure of the integrity of this pathway within cells is the 1α25(OH)₂D₃-mediated induction of 24(OH)ase transcription (8). Our finding that the induction of 24(OH)ase transcription by 1α25(OH)₂D₃ in moDCs was greatest at early time points during DC differentiation, then followed by a significant decline, suggested the possibility that VDR levels change during DC differentiation. To evaluate VDR expression we performed real time RT-PCR from RNA extracts of moDC cultures at a number of time points. The DC-derived amplified product was identical in size to the product from the HL-60 cell line that is known to express VDR (not shown). Immature and mature moDCs demonstrated about a 4-fold reduction in VDR gene transcription as compared with the starting population (Fig. 4⇓A). These changes were mirrored by similar reductions in the level of VDR protein expression as evaluated by nuclear binding of the [³H]1α25(OH)₂D₃ ligand which showed a >2-fold reduction in VDR expression in immature and mature moDCs (Fig. 4⇓B). In contrast to 1αOHase, there was no significant difference between VDR expression (either at the transcript or protein level) in immature and mature DCs. Thus, the down-regulation of VDR was not clearly linked to the terminal differentiation of DCs. Consistent with this interpretation, the decline in VDR transcription was not arrested by specific inhibition of p38 MAPK or NF-κB pathways (data not shown). We also confirmed that neither 1α25(OH)₂D₃ nor 25(OH)D₃ treatment of moDCs was associated with significant differences in VDR expression as compared with controls (data not shown).

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FIGURE 4.

VDR expression is down-regulated during DC differentiation. A, Immature moDCs were propagated as stated previously and matured by CD40 ligation. RNA was extracted at the indicated time points and real time RT-PCR was performed to evaluate VDR expression. Mean fold change (±SEM) in VDR mRNA expression (y-axis) compared with monocytes vs day of culture (x-axis) is shown. Data shown are for eight independent experiments, except for day 3 which represents two independent experiments. Significant differences were evident between day 1 and both days 7 and 9 (p < 0.05 both comparisons). B, VDR expression in moDC cultures was assessed by whole cell nuclear association assays, where monocytes (mo), immature moDCs (imDC), and CD40 ligation-matured moDCs (mDC) were incubated at 1 × 10⁶/ml for 1 h with saturating concentrations of [³H]1α25(OH)₂D₃, washed, and then lysed to form nuclear pellets before scintillation counting. Mean (±SEM) of VDR per cell are shown. Significant differences were evident between day 1 and both days 7 and 9 (p < 0.001 both comparisons). Similar results were obtained when moDCs were matured in the presence of TNF-α. C, moDCs were cultured and immature moDCs were matured by CD40 ligation as before in the presence of vehicle or 1α25(OH)₂D₃ (10⁻⁸ M) added for 72 h from days 1–3, 4–6, or 7–9. Every 72 h cultured cells in all groups were washed extensively and resuspended in fresh media. On day 9 output cells were used a stimulators in an allogeneic mixed leukocyte reaction as outlined in Fig. 2⇑F. The graph shows proliferation (cpm, y-axis) vs number of moDCs per well (x-axis). The result is representative of three independent experiments. Similar results were obtained when moDCs or vitamin D₃ metabolite-treated moDCs were matured in the presence of TNF-α. D, Flow cytometric histograms of CD86 staining in day 9 moDCs/vitamin D₃ metabolite-treated moDCs cultured under the above conditions (isotype control, thin line; day 9 DC, filled histograms). Figures in the upper right corner of each histogram represent mean cellular fluorescence of day 9 DCs stained with specific fluorochrome-labeled Ab minus that of fluorochrome-labeled isotype controls. Results shown are representative of three independent experiments.

The above experiments indicated that as moDCs differentiate, the actions of 1α25(OH)₂D₃ upon DC function could be dependent upon a number of factors, including the expression of VDR, 24(OH)ase activity, and endogenous 1α25(OH)₂D₃ synthesis. Other factors, such as changes in the expression of partner receptors/coactivators to VDR may also modulate the DC response to 1α25(OH)₂D₃. To examine how the net influence of these multiple factors influenced the response of moDCs to 1α25(OH)₂D₃, 10⁻⁸ M 1α25(OH)₂D₃ was added to media on days 1–3, 4–6, or 7–9 following the initiation of moDC culture. After each 72 h, the cells in each culture were washed and resuspended in fresh media with or without the addition of 1α25(OH)₂D₃. As shown in Fig. 4⇑C, the effect of 1α25(OH)₂D₃ ligand upon DC allostimulatory function was reduced the later it was added to developing DC cultures. In parallel, there was a progressive reduction in the inhibitory effect of 1α25(OH)₂D₃ upon expression of cell surface costimulatory molecule expression (Fig. 4⇑D). Thus, as moDCs differentiate in culture, they develop an increasing capacity to synthesize 1α25(OH)₂D₃, but in parallel a reduced ability to respond to its inhibitory actions.