Vitamin D is a steroid hormone that, in addition to its well-characterized role in calcium/phosphate metabolism, has been found to have regulatory properties for immune system function. The nuclear vitamin D receptor is widely expressed in tissues, but has also been shown to be regulated by hormones, growth factors, and cytokines. In this study we show that activation of human Vδ2Vγ9 T cells by nonpeptidic monoalkyl phosphates such as isopentenyl pyrophosphate leads to the up-regulation of the vitamin D receptor via a pathway that involves the classical isoforms of protein kinase C. We further show that this receptor is active by demonstrating that the ligand 1α,25-dihydroxyvitamin D3 (vitD3) significantly inhibits in a dose-dependent fashion phospholigand-induced γδ T cell expansion, IFN-γ production, and CD25 expression. We also show that vitD3 negatively regulates signaling via Akt and ERK and, at high concentrations, potentiates Ag-induced cell death. As such, these data provide further support for the immunoregulatory properties of vitamin D, and suggest that the ability of vitD3 to negatively regulate the proinflammatory activity of γδ T cells may contribute to the protection this vitamin affords against inflammatory and autoimmune disorders dependent upon Th1-type responses.

In this study we examined the potential role of vitamin D in the regulation of T cells expressing the γδ TCR. Vitamin D is a steroid hormone, which in addition to its well-characterized role in calcium/phosphate metabolism and bone maintenance, has been found to have regulatory properties for immune system function (1). The biologically active metabolite 1α,25-dihydroxyvitamin D3 (vitD3)3 appears to function largely, if not exclusively, on a nuclear vitamin D receptor (VDR), which is a member of the superfamily of nuclear steroid, thyroid, and retinoic acid receptors. As such, it acts as a ligand-dependent transcription factor, regulating the activity of target genes that possess DNA sequences containing vitamin D response elements (2, 3). However, nongenomic activity for vitD3 has also been noted, and it remains unclear at the present time whether these effects are also mediated via activation of the nuclear VDR, or whether there is an alternative VDR that is present at the plasma membrane (4). The VDR is widely expressed in tissues, but has also been shown to be regulated at both the transcriptional and posttranslational level by hormones, growth factors, and cytokines (2).

Analyses of tissue distribution of the VDR were the first to suggest that vitD3 might have functions beyond its well-recognized role in calcium and phosphorus metabolism (5), and indeed subsequent studies showed that vitD3 had potent effects on the proliferative activity of several cancerous cell lines, as well as on immune system function in vivo and in vitro (5, 6). In animal models, vitD3 has been shown to have a protective effect against autoimmune diseases such as type 1 diabetes, rheumatoid arthritis (RA), and experimental autoimmune encephalomyelitis (EAE), an animal model of multiple sclerosis (MS). These observations have led to the suggestion that the bioavailability of vitD3, which is predominantly dependent upon exposure to sunlight, could contribute to the increased risk for cancer and diseases such as MS and RA, which have been associated with living at higher latitudes (reviewed in Refs. 6 and 7).

In the immune system, the VDR has been detected in activated T cells (7, 8, 9), and in cells of the monocyte/macrophage series including dendritic cells, whereas expression on B cells remains controversial (7, 10, 11). As might be expected for an essential nutrient, the effect of vitD3 on immune system function depends upon availability and dose. In mice severely depleted of vitD3, blunted T cell responses have been noted, but in normal animals, vitD3 has generally been found to be immunosuppressive. In CD4+ Th cells, vitD3 preferentially inhibits Th1-type cells, particularly the expression of cytokines such as IFN-γ and TNF-α, and to up-regulate the function of Th2 cells by stimulating IL-4 production (12, 13). VitD3 has also been found to inhibit the differentiation and survival of dendritic cells, resulting in impaired alloreactive T cells (14, 15, 16), and to promote the generation of regulatory T cells (CD4+CD25+ T cells) (17).

However, to date, no studies have addressed the effect of vitD3 on γδ T cells. In peripheral blood, γδ T cells represent a minor population of T cells whose exact function remains unknown. T cells that express the Vγ9 (also known as Vγ2)/Vδ2 TCR constitute the major subset and account for 1–10% of the total lymphocyte population in adults (18). In common with other lymphocytes, γδ T cells develop a highly diverse Ag recognition receptor, but the nature of the Ags that activate these cells remain poorly defined. In cells of human origin, the best studied response of these cells is to small phosphate Ags that potently and selectively activate γδ T cells expressing the Vδ2/Vγ9 TCR. These Ags, which are protease resistant but phosphatase sensitive, have been identified as monoalkyl phosphates such as isopentenyl pyrophosphate (IPP) and related prenyl phosphates, which are expressed by a number of potentially pathogenic organisms (19). The response of γδ T cells to these Ags is thought to reflect a role for these cells in host defense via the recognition of families of unprocessed Ags with conserved molecular patterns (20, 21). Activation leads to the rapid production of a number of cytokines, particularly of the Th1 type, and chemokines involved in the initiation of inflammatory and immune responses, as well as in the generation of T cells with cytotoxic activity (22, 23).

In addition to a role in host defense, γδ T cells have also been strongly implicated in the pathogenesis of a number of chronic inflammatory destructive diseases, such as RA and MS, in which Th1-type cytokines and cytotoxic factors are thought to play a critical role. In MS, for example, γδ T cells are present in the inflammatory lesions, and show evidence of clonal expansion (24, 25, 26, 27). Furthermore, in the animal model EAE, depletion of these cells or sensitization of mice with a targeted deletion of the TCR δ locus, results in amelioration of disease expression, at least in the early stages of disease development (28, 29, 30). Because vitD3 has been implicated as a protective factor in MS (31, 32, 33), this raises the question as to whether an effect of vitD3 on the function of γδ T cells could contribute to this result. To address this question, we have examined VDR expression in these cells, and have tested for possible regulatory activity for γδ T cells using the monoalkyl phosphate IPP, or cross-linking through the TCR, as the stimulus for γδ T cell activation. The results show that the VDR is up-regulated in activated γδ T cells and that ligands for the VDR potently modulate the γδ T cell response, implicating vitD3 as an immunoregulatory factor for this specialized subset of T cells.

PBMCs from healthy donors were isolated by Ficoll-Hypaque gradient centrifugation (Pharmacia). Use of human tissues was approved by the Committee on Clinical Investigation of the Albert Einstein College of Medicine. Long-term cultures of Vδ2+ cells were established by stimulating once with IPP (30 μM; Sigma-Aldrich) and were maintained with 50 U/ml IL-2 (National Cancer Institute) as described (34). Vδ2 expression was determined by FACS (clone B6; BD Pharmingen). At the time of testing, Vδ2+ T cells were restimulated with IPP (3 or 10 μM). The biologically active form of vitamin D, vitD3 (Sigma-Aldrich) was dissolved in ethanol and used at concentrations indicated. Control cultures were treated with an equivalent dilution of ethanol. For inhibitor studies, cells were pretreated with the protein kinase C (PKC) inhibitor Gö6976 (250 and 500 nM; Calbiochem), rottlerin (3 and 6 μM; Calbiochem), the MEK1/2 inhibitor UO126 (1–50 μM; Calbiochem), the MEK1 inhibitor PD98059 (25 and 50 μM; New England Biolabs) or the p38 MAPK inhibitor SB203580 (50 μM; Calbiochem).

For analysis of VDR expression, the Abs 9A7 (Biomeda) and D-6 (Santa Cruz Biotechnology) were used. Abs for total and phospho-Akt, and total and phospho-p44/p42 (ERK1/2) were purchased from Cell Signaling Technology. Protein and phosphatase inhibitors were purchased from Sigma-Aldrich. Fluorescein labeled anti-CD25 PE (clone M-A251), anti-IFN-γ PE, anti-IL-10 PE, and anti-Vδ2 FITC or PE were purchased from BD Pharmingen.

Total RNA was collected from γδ T cell control cultures or cultures stimulated with IPP (10 μM) or anti-CD3 Ab (10 μg/ml, HIT3a; BD Pharmingen) for 16 or 24 h using RNeasy Mini kit (Qiagen). Total RNA was amplified using MessageAmp aRNA Kit (Ambion). First-strand cDNAs were synthesized, labeled with Cy5 or Cy3 using a SuperScript II Reverse Transcriptase (Invitrogen) and hybridized to cDNA microarrays containing 27,000 cDNAs representing 18,000 distinct human transcripts. (see 〈http://microarray1k.aecom.yu.edu/〉 for details). Independent measurements of Cy5 and Cy3 signal intensity and background were generated for each cDNA element using a Genepix 4000B scanner (Axon Instruments), and primary data were analyzed using Genepix 3.02 software and normalized by R program (〈www.r-project.org/〉). Experiments for each condition were performed using γδ T cells from at least two different donors.

Vδ2 T cells were washed and adhered to poly-l-lysine-coated slides using a cytospin, fixed with ice-cold methanol for 20 min, washed three times with PBS and blocked in 5% normal goat serum/0.4% Triton X-100/PBS for 20 min at room temperature. Cells were stained for 1 h at room temperature with the D-6 VDR Ab (1:200) in 2% normal goat serum/PBS. After washing twice in PBS, secondary Ab Alexa Fluor 488 goat anti-mouse IgG2a conjugate (Molecular Probes) was applied for 30 min at room temperature, counterstained for 15 min at room temperature with 1 μg/ml 4′,6′-diamidino-2-phenylindole dihydrochloride hydrate (DAPI)/PBS solution and mounted in aqueous mounting medium (gel mount; Biomeda). Fluorescent microscopy was performed on an Olympus IX70 with ×60 numerical aperture 1.4 infinity corrected optics. Images were collected with a Photometrics cooled charge-coupled device camera with a KAF 1400 chip using I.P. Lab Spectrum (Scanalytics).

Generally, γδ T cells were lysed with lysis buffer containing 150 mM NaCl, 0.1% SDS, 50 mM Tris (pH 7.4), 5 mM EDTA, 1% Triton X-100, 1 mM PMSF, 5mM NaF, 5mM Na4P2O7, 1mM Na3VO4, and a mixture of protease inhibitors (Roche Diagnostics). For phosphoERK, after stimulation, cells were directly lysed in 1× Laemmli buffer. Total protein was separated on 10% SDS-PAGE and transferred to polyvinylidene difluoride membrane. They were blocked in TBS/0.1% Tween 20 (TTBS) containing 5% nonfat milk, incubated with primary Abs in 5% BSA/TTBS overnight at 4°C. The blots were washed in TTBS and then further incubated with secondary HRP conjugated Ab (Santa Cruz Biotechnology) in 5% nonfat milk/TTBS for 1 h at room temperature. The reaction was developed using ECL (Pierce) and the blots were quantified using Scion Image.

Freshly isolated PBMC were stimulated with 10 μM IPP at 37°C overnight to induce VDR up-regulation, then restimulated with 10 μM IPP with or without vitD3 (25 or 50 nM) and treated with brefeldin A (10 μM; Sigma-Aldrich). After 6 h at 37°C, the cells were washed twice and stained with anti-Vδ2-FITC in PBS supplemented with 0.5% FCS and 0.01% NaN3. The cells were then fixed with 4% paraformaldehyde, washed twice, and stained intracellularly with anti-IFN-γ-PE, in PBS supplemented with 0.5% saponin (Sigma-Aldrich), 0.5% FCS, and 0.01% NaN3. After 20 min, cells were washed twice with the same solution (0.01% saponin) and analyzed on the MoFlo cytometer (DakoCytomation). Results are expressed as percent of Vδ2+ IFN-γ+ T cells of total Vδ2+ T lymphocytes. Annexin V staining: cells were incubated with annexin V-FITC (BD Pharmingen) and propidium iodide (PI; Sigma-Aldrich) in 100 μl of 1× annexin V binding buffer at room temperature for 15 min. An additional 400 μl of 1× annexin V binding buffer was then added, and cells were analyzed using CellQuest software (BD Biosciences). CD25 expression: cells were double labeled with anti-CD25-PE and anti-Vδ2-FITC on ice for 20 min, washed with PBS, resuspended in 400 μl of PBS, and analyzed using CellQuest software.

Supernatants from Vδ2 T cell lines were harvested at 24 or 48 h poststimulation with IPP (10 μM). Cytokines were quantified by sandwich ELISA per the manufacturer’s instructions. Kits for human IFN-γ and TNF-α were purchased from R&D Systems and for osteopontin from Assay Designs.

Freshly isolated PBMC were stimulated with 10 μM IPP with or without vitD3 (25 or 50 nM). Cells were labeled with CFDA (1 μM; Molecular Probes) and cell divisions followed by flow cytometry. Cells were stained for TCR Vδ2 with anti-Vδ2-PE every day for 1 wk and analyzed on the MoFlo cytometer.

Statistical analysis was performed using Prism software (Prism Software), and data were analyzed using paired student’s t test, assuming an unequal variance with 95% confidence levels, and p values <0.05 were considered significant.

As part of an effort to define the complexity of the pattern of gene expression in γδ T cells activated with phosphate Ags, we used a cDNA transcriptional gene profiling technique comparing the pattern of gene expression in cultures activated with either IPP (10 μM) or Abs to CD3 (10 μg/ml). Activated cells were analyzed against untreated matched controls. Analysis of the results for genes associated with the early activation of T cells, such as Fyn-binding protein, immediate early response 3 gene, and CD25 expression, showed that either IPP or Abs to CD3 activated Vδ2 T cells (Table I). As expected, we also noted potent up-regulation of a number of proinflammatory cytokines and chemokines, many of which have been extensively documented as targets of phosphate-Ag activation in these cells (22, 34) (Table I and data not shown). However, the cDNA screen also showed up-regulation of mRNA for the VDR in cells that had been activated with either IPP or CD3 (Table I). The expression of the VDR in immunocompetent cells has been linked to an immunomodulatory role for vitamin D in the responses of both T cells and dendritic cells (8, 9, 10, 11, 12, 13, 14, 15, 16, 17). Thus, up-regulation of the VDR after restimulation of Vγ9Vδ2 T cells could represent part of a regulatory feedback loop in this population of T cells.

Table I.

Phosphate-containing ligands and anti-CD3 Ab induced gene expression in human Vδ2 T cells in culture

Accession NumberGene NameVδ2
ControlaIPPaFold IncreasebControlaAnti-CD3aFold Increaseb
AA485226 VDR 357 1200 3.4 512 2000 2.7 
  719 6451 9.0 720 2400 4.7 
AA969504 IFN-γ 4500 32000 3600 65000 6.0 
  1500 54478 36 11674 470529 11.0 
NM_000594 TNFLSF2 (TNF-α) 9228 15305 1.7 12519 15883 1.3 
  11483 14450 1.3 16709 18053 1.1 
W72329 TNFLSF1 (lymphotoxin α) 5028 36149 7.2 3839 31408 8.2 
  3809 39144 10.3 7333 27955 3.8 
AA903183 IL-2 receptor, α (CD25) 1447 12481 8.6 1761 8967 5.1 
  1358 19498 14 1382 10309 7.5 
AF198052 FYN-binding protein (FYB-120/130) 1572 7740 4.9 4741 9650 2.0 
  4990 23550 4.7 2513 28215 7.2 
N32077 Immediate early response 3 4196 16436 3.9 1477 8042 5.4 
  2684 23436 8.7 7133 26502 3.7 
Accession NumberGene NameVδ2
ControlaIPPaFold IncreasebControlaAnti-CD3aFold Increaseb
AA485226 VDR 357 1200 3.4 512 2000 2.7 
  719 6451 9.0 720 2400 4.7 
AA969504 IFN-γ 4500 32000 3600 65000 6.0 
  1500 54478 36 11674 470529 11.0 
NM_000594 TNFLSF2 (TNF-α) 9228 15305 1.7 12519 15883 1.3 
  11483 14450 1.3 16709 18053 1.1 
W72329 TNFLSF1 (lymphotoxin α) 5028 36149 7.2 3839 31408 8.2 
  3809 39144 10.3 7333 27955 3.8 
AA903183 IL-2 receptor, α (CD25) 1447 12481 8.6 1761 8967 5.1 
  1358 19498 14 1382 10309 7.5 
AF198052 FYN-binding protein (FYB-120/130) 1572 7740 4.9 4741 9650 2.0 
  4990 23550 4.7 2513 28215 7.2 
N32077 Immediate early response 3 4196 16436 3.9 1477 8042 5.4 
  2684 23436 8.7 7133 26502 3.7 
a

Levels of normalized signal intensities of each of two donors.

b

Fold increase of activated vs control.

We then confirmed the microarray data for the VDR using a combination of Western blotting and immunohistochemistry. Whole cell lysates were prepared from Vδ2+ T cells that had been cultured with IPP for 3 wk and then reactivated with IPP (3, 10, or 20 μM) for varying periods of time. In resting Vδ2+ T cells, no signal was observed on the Western blots (Fig. 1, A and B). However, within 8 h of reactivation with IPP, a clearly detectable band of ∼53 kDa was observed that continued to increase in intensity through 24 h (Fig. 1,A). A more extended period of analysis confirmed prominent induction of the VDR by 24 h with a lower level of expression persisting for several days (Fig. 1,B). Using an Ab specific for the human VDR, immunohistochemistry of control, nonstimulated cells showed only a low level of cytoplasmic immunoreactivity for the VDR (Fig. 1,C). In contrast, cells that had been activated with IPP (3 μM) for 24 h showed robust labeling of T cell aggregates with immunoreactivity for the VDR localized to the nuclear membrane (Fig. 1 C).

FIGURE 1.

Phosphate-containing ligands induce VDR expression in Vδ2 T cells. A, Vδ2 T cell lines were left untreated or activated with IPP (10 μM) for the times shown. The cells were then lysed, separated on 10% SDS-PAGE, and blotted with Ab specific for the VDR. The blot then was stripped and immunoreacted for β-actin as a loading control. One of three separated experiments is shown. B, As in A, but after activation with IPP at 3, 10, or 20 μM for 1–7 days. C, Vδ2 T cell lines were left untreated (left panel) or treated with IPP (10 μM) for 24 h (right panel), and adhered to poly-l-lysine coated slides using a cytospin. Cells were fixed with methanol, stained for VDR expression (green), and counterstained with DAPI (blue) as described in Materials and Methods. Original magnification, ×60.

FIGURE 1.

Phosphate-containing ligands induce VDR expression in Vδ2 T cells. A, Vδ2 T cell lines were left untreated or activated with IPP (10 μM) for the times shown. The cells were then lysed, separated on 10% SDS-PAGE, and blotted with Ab specific for the VDR. The blot then was stripped and immunoreacted for β-actin as a loading control. One of three separated experiments is shown. B, As in A, but after activation with IPP at 3, 10, or 20 μM for 1–7 days. C, Vδ2 T cell lines were left untreated (left panel) or treated with IPP (10 μM) for 24 h (right panel), and adhered to poly-l-lysine coated slides using a cytospin. Cells were fixed with methanol, stained for VDR expression (green), and counterstained with DAPI (blue) as described in Materials and Methods. Original magnification, ×60.

Close modal

The VDR promoter lies in a GC-rich island and does not contain a TATA box, in common with many steroid receptor gene promoters (35). In previous studies, we have shown that culture of Vδ2 cells with IPP results in the activation of a wide-range of signaling pathways, including the classical and novel isoforms of PKC, and the ERK and p38 MAP kinase pathways (34). To assess which of these signaling pathways might be involved in VDR induction, we used specific inhibitors in Vδ2+ T cells activated with IPP. We also tested an inhibitor of PKA, because this signaling pathway has been implicated in VDR up-regulation in osteoclasts (36). As shown in Fig. 2, the PKA inhibitor H-89 had no effect, whereas the inhibitor of the classical PKC pathway GÖ6976 potently inhibited IPP-induced VDR expression in a dose-dependent fashion (Fig. 2,B). The putative novel PKC inhibitor rottlerin, which we have shown previously can inhibit IPP-induced AP-1 and NF-κB activation in γδ T cells (34), was not as potent (Fig. 2,B). We next examined the effect of inhibiting ERK activation by applying the MEK inhibitors UO126 and PD98059. Both UO126 and PD98059 blocked IPP-induced VDR expression as determined by Western blotting, although UO126 was more potent (Fig. 2,C). UO126 has been shown to inhibit the activated phosphorylated form of MEK1/2, whereas PD98059 appears to primarily inhibit MEK1/2 activation by blocking the access of activating enzymes (37). The p38 MAP kinase inhibitor SB203580 had only a minimal effect. Densitometric analysis of data shown in Fig. 2, AC, expressed as percent of VDR expression from that detected in the untreated controls, after normalization, is shown in Fig. 2 D.

FIGURE 2.

Signaling pathways involved in IPP-induced VDR expression. A, Vδ2 T cell lines were left untreated or treated with IPP (3 μM) for 24 h in the absence or presence of 40 or 80 nM of the PKA inhibitor H-89. B, As in A, but using the classical PKC inhibitor Gö6976 (250 or 500 nM) or the putative novel PKC inhibitor Rottlerin (3 or 6 μM). C, As in A, but using the ERK signaling pathway inhibitors UO126 (1, 5, or 25 μM) and PD98059 (25 or 50 μM), or the P38 inhibitor SB203580 (50 μM). D, Densitometric analysis of data shown in A–C expressed as percent VDR expression level in the untreated controls after normalization.

FIGURE 2.

Signaling pathways involved in IPP-induced VDR expression. A, Vδ2 T cell lines were left untreated or treated with IPP (3 μM) for 24 h in the absence or presence of 40 or 80 nM of the PKA inhibitor H-89. B, As in A, but using the classical PKC inhibitor Gö6976 (250 or 500 nM) or the putative novel PKC inhibitor Rottlerin (3 or 6 μM). C, As in A, but using the ERK signaling pathway inhibitors UO126 (1, 5, or 25 μM) and PD98059 (25 or 50 μM), or the P38 inhibitor SB203580 (50 μM). D, Densitometric analysis of data shown in A–C expressed as percent VDR expression level in the untreated controls after normalization.

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To determine whether ligation of the VDR can inhibit IPP-induced activation of γδ T cells, we first assessed the effect of the activated form of vitD3 on IFN-γ production that we, and others, have shown is potently stimulated by IPP in γδ T cells (Refs. 22 , 23 , and 34 ; see also Table I). We used two approaches to address this question. In the first, freshly isolated PBMC were cultured overnight with IPP (10 μM) to stimulate VDR expression. Cells were restimulated with IPP (10 μM) without and with the addition of vitD3 (25 or 50 nM) and cultured for 6 h. IFN-γ production was then determined by intracellular staining. No IFN-γ was detected in resting cells, however after stimulation with IPP ∼40% of γδ T cells expressed IFN-γ intracellularly (Fig. 3 A). A small increase was also noted in non-γδ T cells. Culture with vitD3 (25 nM) markedly inhibited IPP-induced IFN-γ production in both populations of T cells. Cells cultured with vitD3 (25 or 50 nM) in the absent of IPP had no effect (data not shown). Cumulative data from three different donors showed a 37 ± 10% reduction of IFN-γ production for 25 nM vitD3 or a 53 ± 18% reduction for 50 nM vitD3.

FIGURE 3.

VitD3 regulates IPP-induced IFN-γ expression. A, Freshly isolated PBMCs were activated with IPP (10 μM) in the absence or presence of vitD3 (25 or 50 nM) overnight. The cells were then stained for intracellular IFN-γ as described in Materials and Methods and analyzed by FACS. The percent of IFN-γ-positive cells is shown in the upper right quadrant. B, Vδ2 T cell lines were treated and processed as in A. C, Cumulative FACS data for two Vδ2 T cell lines derived from two different donors are shown. D, Vδ2 T cell lines were activated with IPP (10 μM) plus ethanol as a diluent control or with vitD3 at 50, 25, or 10 nM, and the supernatants were collected at 24 and 48 h. IFN-γ levels were determined by ELISA (n = 2). No IFN-γ was detected in cell supernatants taken from unstimulated cells (data not shown). E, As in D, except that cells were preincubated with IPP (10 μM) for 24 h before the addition of vitD3 at 100, 50, or 25 nM (n = 2).

FIGURE 3.

VitD3 regulates IPP-induced IFN-γ expression. A, Freshly isolated PBMCs were activated with IPP (10 μM) in the absence or presence of vitD3 (25 or 50 nM) overnight. The cells were then stained for intracellular IFN-γ as described in Materials and Methods and analyzed by FACS. The percent of IFN-γ-positive cells is shown in the upper right quadrant. B, Vδ2 T cell lines were treated and processed as in A. C, Cumulative FACS data for two Vδ2 T cell lines derived from two different donors are shown. D, Vδ2 T cell lines were activated with IPP (10 μM) plus ethanol as a diluent control or with vitD3 at 50, 25, or 10 nM, and the supernatants were collected at 24 and 48 h. IFN-γ levels were determined by ELISA (n = 2). No IFN-γ was detected in cell supernatants taken from unstimulated cells (data not shown). E, As in D, except that cells were preincubated with IPP (10 μM) for 24 h before the addition of vitD3 at 100, 50, or 25 nM (n = 2).

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In the second approach, we prepared γδ T cell lines and tested for IPP-induced IFN-γ by both FACS analysis and ELISA. In nonstimulated Vδ2+ cell, no IFN-γ was detected either by FACS analysis (Fig. 3,B) or by ELISA (data not shown). However, after activation with IPP (10 μM) for 16 h, strong intracellular staining for IFN-γ was noted in ∼29 ± 8% of the total γδ T cell population. The addition of vitD3 (25 nM) to the medium at the same time as IPP resulted in a 42 ± 14% reduction in the number of cells staining positively for IFN-γ. Cells cultured with vitD3 (25 nM) in the absence of IPP had no effect (representative donor shown in Fig. 3,B). Pooled data for two separate donors is shown in Fig. 3,C. To provide a more quantitative assessment of the effect of vitD3 on IFN-γ production, we quantified IFN-γ in the cell supernatants by ELISA using two different protocols. In the first, vitD3 was added simultaneously with IPP, and supernatants were assayed 24 and 48 h posttreatment (Fig. 3,D). In the second, vitD3 was added 24 h poststimulation with IPP, at a time when the VDR expression would be expected to be maximal, and supernatants were harvested at 24 and 48 h postaddition of vitD3 (Fig. 3 E). Control cultures were treated with ethanol alone. The results showed that vitD3 dose-dependently reduced IFN-γ production using both protocols, but a more potent inhibition of IFN-γ release was obtained when vitD3 was applied 24 h post-IPP stimulation.

In other PBMC populations, vitD3 has been shown to promote the production of IL-10 (17) and osteopontin (38). However, ELISA analysis of supernatants derived from control and IPP-stimulated γδ T cells, as well as intracellular staining for IL-10, failed to detect the presence of these cytokines in these cells (data not shown).

To explore further the effect of vitD3 on γδ T cell activation, we studied expression and function of CD25, the high affinity IL-2R. As shown in Table I, activation with IPP results in the induction of CD25 expression (34), and FACS analysis confirmed this result, showing that in control resting Vδ2+ cells, CD25 was not expressed but was strongly up-regulated by IPP (Fig. 4,A). The addition of vitD3 reduced this expression by ∼25%. Cumulative data with cells derived from four different donors showed that vitD3 statistically significantly reduced CD25 expression on cells stimulated with IPP plus vitD3 compared with cells stimulated with IPP alone (Fig. 4 B).

FIGURE 4.

VitD3 regulation of IPP-induced CD25 expression and function. A, Vδ2 T cell lines were activated with IPP (10 μM) in the absence or presence of vitD3 (25 nM) and CD25 on the cell membrane assessed by FACS. B, Cumulative FACS data for four different vδ2 T cell lines are shown. Asterisk denotes values significantly different from the IPP treated culture: ∗, p < 0.01. C, Vδ2 T cells were stimulated overnight with IPP to induce expression of both CD25 and the VDR and then rested in medium without IL-2 for 4 h. They were then stimulated with IL-2 (50 U/ml) with or without vitD3 (25 nM) for the times shown, and phosphorylation of Akt and ERK was determined by Western blotting.

FIGURE 4.

VitD3 regulation of IPP-induced CD25 expression and function. A, Vδ2 T cell lines were activated with IPP (10 μM) in the absence or presence of vitD3 (25 nM) and CD25 on the cell membrane assessed by FACS. B, Cumulative FACS data for four different vδ2 T cell lines are shown. Asterisk denotes values significantly different from the IPP treated culture: ∗, p < 0.01. C, Vδ2 T cells were stimulated overnight with IPP to induce expression of both CD25 and the VDR and then rested in medium without IL-2 for 4 h. They were then stimulated with IL-2 (50 U/ml) with or without vitD3 (25 nM) for the times shown, and phosphorylation of Akt and ERK was determined by Western blotting.

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We next determined whether signaling through the IL-2 receptor was directly affected by vitD3. To address this question, cells were stimulated overnight with IPP to induce expression of both CD25 and the VDR. Cells were rested in medium without IL-2 for 4 h, and then treated with IL-2 for the times shown. Ligand-induced activation was determined by Western blotting for phosphorylation of Akt and ERK. As shown in Fig. 4 C, the addition of IL-2 to the medium resulted in potent phosphorylation of both Akt and ERK. The presence of vitD3 (25 nM) led to both a reduction in the extent of phosphorylation of Akt and ERK, as well as a reduction in the persistence of the phosphorylated state. In contrast, vitD3 addition in the absence of IPP had no effect on either Akt or ERK.

In other cell types, vitD3 has been shown to inhibit proliferation (39). To test for this in IPP-activated γδ T cells, we labeled freshly isolated PBMC with CFDA and monitored cell proliferation by FACS over a 7-day period. Cells were stimulated with IPP without or with different doses of vitD3 (10, 25, or 50 nM). As shown in Fig. 5,A, vitD3 dose-dependently reduced the percentage of γδ T cells detected in these cultures after activation with IPP, whereas incubation with IL-2 alone (control) or vitD3 (25 nM) alone had no effect on the survival/proliferation of γδ T cells. Cumulative data for three or four donors for these experiments with different concentrations of vitD3 (25 or 50 nM) are shown in Fig. 5,B. The ERK family of serine/threonine kinases plays a pivotal role in intracellular signaling pathways involved in proliferation, and thus we examined whether the regulatory effect of vitD3 for IPP-induced proliferation was reflected in an effect on ERK phosphorylation. As shown in Fig. 5,C, reactivation with IPP led to a prolonged increase in phosphorylation of both ERK1 (44 kDa) and ERK2 (42 kDa), and this was markedly reduced by the addition of vitD3 (25 nM). Densitometric analysis of the blots is shown in Fig. 5 D.

FIGURE 5.

The effect of VitD3 on IPP-induced proliferation and cell death. A, Freshly isolated PBMCs were activated with IPP (10 μM) in the absence or presence of vitD3 (25 nM). Cells were loaded with CFDA, and fluorescence intensity was determined by FACS. Results are shown for 5 and 7 days post-IPP stimulation. Boxed areas represent Vδ2 T cells. B, Data collected as in A from three or four different donors. Percentage of Vδ2 T cells was determined using gates as shown in A. On day 7, the values between the control IPP-treated cells and the IPP+vitD3 at 25 or 50 nM were statistically significantly different, p < 0.03. C, Vd2 T cells were stimulated with IPP (10 μM) overnight to induce the expression of VDR, then restimulated with IPP (10 μM) with or without vitD3 (25 nM) for 30 or 60 min, and phospho-ERK and total ERK were determined by Western blotting. D, Densitometric analysis of data shown in C expressed as ratio of phospho-ERK to total ERK. E, Vδ2 T cell lines were treated overnight with IPP (10 μM) in the absence or presence of VitD3 at 50 or 100 nM. Then cells were stained with FITC-conjugated anti-Vδ2TCR (FL1). Cells were analyzed by FACS, and two populations (gates R1 and R2) were defined by size and intensity of TCR Vδ2 fluorescence. F, Cumulative data for three donors treated with 100 nM vitD3 and analyzed as in E. G, Representative FACS analysis of one Vδ2 T cell line stained for annexin V and PI as described in Materials and Methods. H, Cumulative FACS data for three different Vδ2 T cell lines stained for annexin V and PI. Asterisk denotes values significantly different from the IPP-treated culture: ∗, p < 0.001.

FIGURE 5.

The effect of VitD3 on IPP-induced proliferation and cell death. A, Freshly isolated PBMCs were activated with IPP (10 μM) in the absence or presence of vitD3 (25 nM). Cells were loaded with CFDA, and fluorescence intensity was determined by FACS. Results are shown for 5 and 7 days post-IPP stimulation. Boxed areas represent Vδ2 T cells. B, Data collected as in A from three or four different donors. Percentage of Vδ2 T cells was determined using gates as shown in A. On day 7, the values between the control IPP-treated cells and the IPP+vitD3 at 25 or 50 nM were statistically significantly different, p < 0.03. C, Vd2 T cells were stimulated with IPP (10 μM) overnight to induce the expression of VDR, then restimulated with IPP (10 μM) with or without vitD3 (25 nM) for 30 or 60 min, and phospho-ERK and total ERK were determined by Western blotting. D, Densitometric analysis of data shown in C expressed as ratio of phospho-ERK to total ERK. E, Vδ2 T cell lines were treated overnight with IPP (10 μM) in the absence or presence of VitD3 at 50 or 100 nM. Then cells were stained with FITC-conjugated anti-Vδ2TCR (FL1). Cells were analyzed by FACS, and two populations (gates R1 and R2) were defined by size and intensity of TCR Vδ2 fluorescence. F, Cumulative data for three donors treated with 100 nM vitD3 and analyzed as in E. G, Representative FACS analysis of one Vδ2 T cell line stained for annexin V and PI as described in Materials and Methods. H, Cumulative FACS data for three different Vδ2 T cell lines stained for annexin V and PI. Asterisk denotes values significantly different from the IPP-treated culture: ∗, p < 0.001.

Close modal

The reduced expansion of IPP-activated γδ T cells in the presence of vitD3 shown above could result from either a delay in proliferation or a cytotoxic effect. To determine whether vitD3 was toxic for γδ T cells, we assessed evidence of cell death in γδ T cell lines by estimating changes in cell size as well as staining for annexin V and PI as markers of apoptosis (40). As shown in Fig. 5,E, addition of IPP (10 μM) overnight led to the accumulation of a distinct population of γδ T cells with a smaller profile (defined by the R2 gate). Costimulation with IPP plus vitD3 at 25 (data not shown) or 50 nM did not alter the ratio of cells in the R1 and R2 gates either in the control cultures or in IPP-activated cultures (Fig. 5,E, upper panel). However, after addition of 100 nM vitD3, an increase in the population of cells in the R2 gate was noted (Fig. 5,E, lower panel). Quantitative analysis for three donors is shown in Fig. 5,F. The extent of apoptosis in the R1 and R2 populations of cells was then determined using FACS analysis of annexin V and PI staining (Fig. 5, G and H). As shown in Fig. 5,G, in control resting Vδ2+ T cells only a few cells displayed evidence of apoptosis (4%) and addition of 100 nM vitD3 resulted in only a slight increase in this number (7%). However, after activation with IPP, a dramatic increase was noted in the population that was double-stained for both annexin V and PI (20%). This was further increased in cell cultures to which vitD3 (100 nM) had been added (35%). This resulted in an overall reduction in the viable cell population. Further analysis showed that these apoptotic cells were present only within the R2 gate (data not shown). This experiment was then repeated with cells from an additional two donors, and the combined data for these three donors is presented graphically in Fig. 5 H.

In this study we have shown that expression of the VDR can be up-regulated in γδ T cells after activation by either phosphate ligands or cross-linking of the TCR through a pathway that involved activation of the classical isoforms of PKC. We have also shown that this receptor is functional by demonstrating that the ligand vitD3 negatively regulated IFN-γ production, inhibited phosphate-Ag-induced γδ T cell activation as determined by reduced expression of CD25, and inhibited Ag-induced expansion in a dose-dependent fashion. At high concentrations, vitD3 also potentiated Ag-induced cell death. As such, these activities of vitD3 would be expected to result in a general dampening of the stimulatory properties of phosphate Ags for this minor subset of T cells.

The observation that VDR expression is up-regulated via a mechanism that involves PKC is in agreement with previous studies using rat osteosarcoma cell (41) and LLCPK-1 porcine kidney cells (42). Additional studies that identified a functional AP-1 site in the mouse VDR promoter (43), and that detected increased expression of the human VDR in LLCPK-1 cells cotransfected with a PKCα expression vector (41), would also be consistent with a role for PKC as a positive regulator of the VDR. However, additional reports using UMR-106 osteoclasts, rat intestinal epithelial cells, and mouse fibroblasts detected either no change or a decrease in VDR levels after PKC activation (44, 45, 46). Thus, the role of PKC in VDR up-regulation appears to be cell-type specific. Because both signaling via the TCR and PKC can also lead to MEK-dependent ERK phosphorylation, we also determined whether inhibition of MEK altered IPP-induced VDR expression. The data showed that the MEK inhibitors U-0126 and PD-98059 only partially blocked phosphate-Ag-induced VDR expression, suggesting that both MAPK-dependent and -independent pathways are involved in VDR expression in γδ T cells.

The activation of γδ T cells by phosphate Ags has been shown to induce the release of a wide range of cytokines and chemokines with known proinflammatory and Th1-like activities (22, 23, 34). VitD3 has been shown to regulate both negatively and/or positively several cytokines including IFN-γ, IL-2, IL-12, TNF, IL-10, osteopontin, and GM-CSF (10, 11, 12, 13, 14, 15, 16, 17). γδ T cells are a poor source of IL-2 and IL-10, and we could not detect osteopontin in these cells after activation with phosphate Ags either by microarray or ELISA (data not shown). Thus, we tested for an effect of vitD3 on phosphate-Ag-induced IFN-γ. Our data showing that vitD3 led to decreased expression of IFN-γ in γδ T cells is in agreement with several previous studies both in vitro and in vivo that have documented a role for this receptor in the negative regulation of IFN-γ expression in other lymphocyte populations (47, 48). CD4 T cells derived from VDR knockout mice have also been found to express increased levels of IFN-γ (49). That this effect of IFN-γ expression likely reflects a direct effect on gene expression has come from studies in Jurkat cells, where it has been shown that VDR-mediated repression involved the formation of a heterodimer between the VDR and the retinoid-X-receptor (VDR-RXRα), which bound to two different sites on the IFN-γ promoter (48). In our studies, exposure of cells to vitD3 either at the time of activation with phosphate Ags, or 24 h postactivation when the VDR levels were found to be maximally elevated, effectively suppressed IFN-γ expression. This would suggest that even though the VDR is an inducible protein in γδ T cells, ligands for this receptor could target cells undergoing Ag-induced stimulation as well as cells that are fully activated at sites of inflammation.

The generally down-regulatory effect of vitD3 for Th1-type cytokines, along with the positive regulatory effects for the Th2-type cytokine IL-4 and the Th2 transcription factor GATA3 (12), has been implicated as a contributing factor for the suppressive effect of vitD3 for Th1-type diseases such as EAE (50, 51, 52), experimental arthritis (53), and experimental inflammatory bowel disease (54). An alternative, but not necessarily exclusionary, hypothesis for the beneficial effects of vitD3 for these Th1-like diseases is that it reflects the activity of vitD3 on dendritic cells. In dendritic cells, vitD3 potently inhibits the expression of multiple maturation-induced proteins (16), as well as Ag-presenting activity, resulting in the development of tolerized T cells, or T cells with regulatory activity (17).

In follow-up studies that have addressed the mechanism of action of vitD3 in the passive-transfer model of EAE, Nashold et al. (55, 56) concluded that the data strongly implicated the requirement for the generation of a Rag-1-dependent cell that specifically limited the occurrence of activated, autoreactive T cells in the CNS, supporting the hypothesis that vitD3 is required for the optimal functioning of regulatory T cells that maintain self-tolerance in the periphery. However, γδ T cell depletion has also been shown to protect against EAE (28, 29, 30) and in these models it has been proposed that the proinflammatory activities of γδ T cells may act to lower the threshold for Th1-type T cell activation, thus facilitating the accumulation of Ag-specific T cells in the CNS. Therefore, it could be argued that vitD3 may act not only to promote a negatively acting regulatory T cell (17), but also to inhibit a positively acting regulatory T cell, perhaps expressing the γδ rather than the αβ TCR. Additional studies from Hayes and colleagues (57) have suggested that vitD3 also reverses EAE by stimulating inflammatory cell apoptosis. Our data showing that high-dose vitD3 augmented Ag-induced cell death, while not showing toxicity in the absence of Ag, lends support to this concept, and further suggest that this may be mediated by a reduction in CD25 expression, because apoptotic cells were found exclusively in the population of cells expressing low levels of CD25, the high-affinity IL-2 receptor (data not shown). IL-2 is a required growth factor for γδ T cells, and IL-2 signaling was also rapidly down-regulated by vitD3 through the Akt pathway, a signaling pathway known to be involved in cell survival. However, it is important to note that in T cells in which Ag-induced cell death proceeds via a Fas-mediated pathway, vitD3 has been found to suppress apoptosis through an interaction of activated VDR with a vitamin D response element in the fasL gene, resulting in repression of ligand-dependent transcriptional activation (58). Thus, vitD3 may also regulate T cell responses by leading to the selective elimination/survival of specific subpopulations of lymphocytes. We have shown recently that at least two distinct subsets of Vδ2+ T cells can be defined in PBMCs, and studies are ongoing to determine whether vitD3 has a differential effect on these supopulations of Vδ2+ T cells (59).

In conclusion, our data support a role for vitD3 in the regulation of the γδ T cell response to phosphate Ags and lend further support to the concept that up-regulation of the VDR, in conjunction with ligand activation, forms part of negative feedback loop at sites of inflammation. This would be consistent with data showing a protective effect of γδ T cell depletion in several inflammatory animal models such as EAE (28, 29, 30). However, it is important to note that the activation of γδ T cells by phosphate Ags has been linked to a role for these cells in protection against bacterial, particularly mycobacterial, infections (18). Poor vitamin D nutrition, as well as certain alleles of the VDR, correlates with susceptibility to mycobacterial infections. These observations have led to the use of vitamin D as a vaccine adjuvant, where it was found that vitamin D enhanced mucosal immunity, further supporting a role for this pathway in boosting Th2-type responses. Perhaps of even greater interest are the recent data that show that vitamin D response element are present in the promoters of several antibacterial peptides including the human cathelicidin antimicrobial peptide (camp) and defensin β2 (defB2) (60). Moreover, vitD3 was found to induce increases in the antimicrobial activity of human monocytes and neutrophils via an increase in antimicrobial proteins. Thus, the possibility that vitD3 may lead to the selective down-regulation of the inflammatory properties of γδ T cell while enhancing the antibacterial properties of these cells will be of interest to determine in future studies.

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 U.S. Public Health Service Grants NS 31919 and NS 11920 and Institutional Grants from the Italian Ministry of Health.

3

Abbreviations used in this paper: vitD3, 1α,25-dihydroxyvitamin D3; VDR, vitamin D receptor; EAE, experimental autoimmune encephalomyelitis; MS, multiple sclerosis; IPP, isopentenyl pyrophosphate; CFDA, carboxyfluorescein diacetate; PI, propidium iodide; PKC, protein kinase C; RA, rheumatoid arthritis.

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