MC-T_reg OX40-mediated interaction inhibits Ca²⁺ influx independent of PLC-γ activation or intracellular Ca²⁺ mobilization

To explore the underlying mechanism for the inhibitory effect of T_regs on MCs degranulation, we investigated signaling events known to be essential for MCs degranulation. Very little is known about the signals generated subsequent to OX40L stimulation, but it has been published that OX40L engagement results in the rapid translocation of the Ca²⁺-dependent protein kinase C (PKC) β to the membrane of human airway smooth muscle cells (Burgess et al., 2004). Ca²⁺ mobilization and PKCβ activation are known to be absolutely essential for MCs degranulation (Blank and Rivera, 2004). Ca²⁺ mobilization is initiated by the phosphorylation of PLC-γ following FcεRI engagement, which leads to the hydrolysis of phosphatidylinositol-4,5-biphosphate to inositol 1,4,5-triphosphate (IP₃) and diacylglycerol (DAG). IP₃ causes mobilization of intracellular Ca²⁺ ([Ca²⁺]_i) from endoplasmic reticular stores, whereas DAG and Ca²⁺ act in concert to promote activation of Ca²⁺-dependent PKCs, like PKC-β. The emptying of internal stores triggers Ca²⁺ influx from external sources, a step required for MCs degranulation (Gilfillan and Tkaczyk, 2006).

We first explored whether a defect in the activation of PLC-γ2 (as measured by phosphorylation at Y759 site, known to be required for lipase activity) was defective. The −γ2 isoform was chosen for its essential role in MCs degranulation (Wen et al., 2002). Rapid tyrosine phosphorylation of the PLC-γ2 isoform was detected upon FcεRI engagement in both WT and OX40L^−/− BMMCs (Figure 3A, B and C). While a trend towards a slightly more transient phosphorylation was observed in OX40L^−/− BMMCs, this was not significant. Moreover, the presence of WT or OX40^−/− T_regs did not significantly alter the phosphorylation of PLC-γ2 in either WT or OX40L^−/− BMMCs (Figure 3D and E).

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Figure 3

T_regs do not affect FcεRI-dependent PLC-γ phosphorylation

WT or OX40L^−/− sensitized BMMCs were stimulated with Ag in the presence of WT or OX40^−/− T_regs for the indicated times. Cells were immediately fixed and stained for c-kit and phosphorylated PLC-γ2. From c-kit⁺-gated BMMCs (A), histogram overlays of phosphorylated PLC-γ2 at different time points were obtained from WT (upper panels) or OX40L^−/− (lower panels) BMMCs challenged in absence of T_regs (B). Dot plot overlay of basal phosphorylated PLC-γ2 (left, gray) and after 10 min (right, violet) is shown in panel C. Histogram overlays of phosphorylated PLC-γ2 from WT (upper panels) or OX40L^−/− (lower panels) BMMCs challenged in the presence of T_regs (D). Results shown are representative of three independent experiments. Kinetics of PLC-γ2 phosphorylation at different conditions are shown in panel E and are the mean + SD of three independent experiments.

Recent studies demonstrated that in MCs extracellular Ca²⁺ influx can be regulated independently of PLC-γ, for example through the activity of sphingosine kinase 2 (Olivera et al., 2007). Thus, investigating the effects of WT and OX40^−/− T_regs on the Ca²⁺ response of MCs was warranted. Interestingly, we observed that FcεRI-dependent Ca²⁺ mobilization in MCs is impaired in the presence of WT but not OX40^−/− T_regs (Figure 4A). As PLC-γ2 activation was largely unaffected, this suggested that the effect on Ca²⁺ mobilization was likely independent from the emptying of intracellular stores. Indeed, the depletion of extracellular Ca²⁺ showed that MCs had a relatively normal mobilization of Ca²⁺ from intracellular stores in the presence of T_regs. However, restoration of Ca²⁺ in the extracellular medium revealed a considerable defect in Ca²⁺ influx (Figure 4B). The ability or inability of these MCs to flux Ca²⁺ across the plasma membrane in the absence or presence of T_regs was consistent with their normal or decreased MCs degranulation, respectively (Figure 4C and D).

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Figure 4

Reduced FcεRI-dependent Ca²⁺ influx following BMMC-T_reg engagement, but not intracellular Ca²⁺ mobilization

(A) BMMCs loaded with FURA-2AM were stimulated via FcεRI in the absence (BMMC, black line) or presence of CD4⁺CD25⁻ T cells (T resting, green line), WT CD4⁺CD25⁺ T_regs (red line) or OX40^−/− CD4⁺CD25⁺ T_regs (blue line) and fluorescence emission was monitored. (B) FURA-2AM-loaded BMMCs were stimulated via FcεRI and co-cultured with WT CD4⁺CD25⁺ T_regs (red line) in the absence of extracellular Ca²⁺. 400 sec after Ag stimulation, 2μM Ca²⁺ was added to the medium and fluorescence emission was monitored. (C and D) At the end of each experiment, 14 minutes after Ag addition, percentage of β-hexosaminidase release from individual sample, was measured. Results shown are representative of three independent experiments.

cAMP is involved in T_regs-mediated suppression of BMMCs activation

Our data indicated that T_regs directly inhibited MC degranulation by cell-cell contact, through the interaction of OX40 on T_regs with OX40L on MCs, resulting in the suppression of Ca²⁺ influx in MCs upon FcεRI stimulation. However, how these events are linked remained to be explored. T_regs have been reported to block effector T cell functions by producing cAMP. Thus, one possibility was that T_reg-mediated suppression of MCs might occur through T_reg production of cAMP, which could lead to decreased Ca²⁺ influx and suppression of degranulation in BMMCs, as recently shown (Hua et al., 2007). We first confirmed this effect by treating MCs with forskolin, which raises the intracellular cAMP concentration, and found that this treatment caused inhibition of MCs degranulation (Supplemental, Figure S4) and induced a 1.5 fold increase in cAMP levels in BMMCs (Figure 5A). When IgE/Ag-activated BMMCs were incubated in the presence of T_regs, cAMP levels in sorted BMMCs increased by approximately 3 fold (Figure 5A). No increase in cAMP was observed when BMMCs were co-cultured with OX40^−/− T_regs or when OX40L^−/− BMMC were co-cultured with T_regs (Figure 5A). No significant differences were found in the intracellular levels of cAMP in WT and OX40^−/− T_regs alone or co-incubated with BMMCs (Figure 5B). Therefore, these findings showed that constitutive OX40-OX40L interaction between T_regs and MCs resulted in significant increase of cAMP only in the MCs. To address the possibility that OX40-OX40L interactions might enhance gap junction formation, resulting in cAMP transfer from T_regs to MCs as previously described for T_reg-T_effector interactions (Bopp et al., 2007), we first incubated BMMCs with OX40-expressing membranes from the K562 cell line and found cAMP increase MCs, albeit at lower levels than with intact cells (Figure 5C). This was consistent with the more modest inhibitory effect of such OX40-expressing membranes on MC degranulation (Figure 2C), which likely reflects a less extensive engagement of OX40L by K562 OX40-expressing membranes relative to intact T_regs. To fully exclude the passage of cAMP from T_regs to MCs, we measured the transfer of calcein, a dye that diffuses only via gap junction (Fonseca et al., 2006), and found no transfer (Supplemental, Figure S5). These data suggest that the rise of cAMP in MCs is likely to result from intracellular signals induced by T_regs to MCs through OX40L engagement. To determine if the reversal of cAMP increase would cause reversal of the inhibitory effects, we pretreated MCs with the antagonist Rp-cAMP (shown to block cAMP-dependent PKA activity) and then tested MCs degranulation and Ca²⁺ mobilization in presence of T_regs. Treatment with Rp-cAMP did not alter either cell viability or the threshold of MCs activation (data not shown). Rp-cAMP treated BMMCs were resistant to the inhibitory effects of T_regs upon FcεRI stimulation, showing a degranulation response identical to FcεRI-stimulated MCs in the absence of T_regs (Figure 6A). Moreover, in presence of the Rp-cAMP the uptake of extracellular Ca²⁺ was unaffected by the presence of T_regs (Figure 6B).

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Figure 5

Intracellular levels of cAMP are increased in BMMCs after co-culture with WT but not with OX40-deficient T_regs

Sensitized WT and OX40L^−/− BMMCs were CFSE-labeled and Ag-stimulated alone or with WT or OX40^−/− T_regs. BMMCs and T_regs were separated using FACS-based cell sorting and cytosolic cAMP concentrations were measured using a cAMP-specific ELISA. As positive control sensitized BMMCs were incubated with forskolin and challenged with Ag. (A) BMMCs baseline [cAMP] was 10 pmoles/1×10⁵. Results are expressed as fold induction over BMMCs alone. (B) T_regs baseline [cAMP] was 50 pmoles/1×10⁵. Results are expressed as fold induction over T_regs alone. (C) Sensitized BMMCs were activated with Ag plus K562 or K562-OX40 membranes. The mean ± SD of three independent experiments are shown.

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Figure 6

Antagonism of cAMP effects in BMMCs reverses T_reg-mediated suppression of degranulation and restores extracellular Ca²⁺ uptake

(A) Anti-DNP IgE preloaded BMMCs were preincubated for 30 min with 1mM of the specific cAMP antagonist Rp-cAMPs. Cells were washed and activated with Ag separately (BMMC) or in co-culture with CD4⁺CD25⁺ T_regs (BMMC + T_reg). After 30 min samples were examined for release of β-hexosaminidase as described. Shown are the means ± SD of three independent experiments each performed in duplicate. (B) IgE-sensitized BMMCs were preincubated for 30 min with 1mM of the specific cAMP antagonist Rp-cAMPs and Ca2+ mobilization was assessed.

Lymph node immunolabeling

For double immunohistochemical staining, 4μm-thick sections were cut from formalin-fixed paraffin-embedded inguinal lymph node samples from 8 weeks old C57BL/6. Slides were pre-incubated with protein block (Novocastra, UK), incubated with mouse anti-rat-FcεR (clone IRK) (Rivera et al., 1988) followed by biotinilated swine anti-mouse Ab, streptavidin-conjugated alkaline phosphatase (LSAB+ kit, Dako, Denmark) and labeled using the Fast Red chromogenic substrate (Dako). Sections were incubated with the primary rat anti-mouse-Foxp3 Ab (clone FJK-16s, eBiosciences, San Diego, California), secondary horseradish peroxidase (HRP)-conjugated anti-rat-Ig and labeled with hydrogen peroxide/diaminobenzidine (DAB+) (Dako). Imunohistochemical evaluation was performed using a Leica DM2000 optical microscope (Leica Microsystems, Germany) and microphotographs were collected using Leica DFC320 digital camera (Leica Microsystems).

Purification of CD4⁺CD25⁺ and CD4⁺CD25⁻ subsets

CD4⁺CD25⁺ cells were purified using the CD25⁺ T-cell isolation kit (Miltenyi Biotec, Bergisch Gladbach, Germany) according to manufacturer’s instructions. Flow cytometry showed that the separated fractions were more than 90% Foxp3⁺(Supplemental, Figure S2). For some experiments CD4⁺CD25⁻ T cells were stimulated 72 h with 1 μg/mL of plate-coated anti-CD3 plus 2.5 μg/mL of soluble anti-CD28 (both from eBioscience).

Bone marrow-derived mast cell differentiation, activation and FcεRI expression

BMMCs were obtained by in vitro differentiation of bone marrow cells taken from mouse femur as described (Frossi et al., 2007). After 5 weeks, BMMCs were monitored for FcεRI expression by flow cytometry. Purity was usually more than 97%. BMMCs were obtained from three to four mice and all experiments are performed using at least three individual BMMC cultures. Before experiments, 1 × 10⁶/ml BMMCs were sensitized in medium without IL-3 for 4 hr with 1μg/ml of DNP-specific IgE and challenged with DNP-HSA in Tyrode’s buffer (10 mM HEPES buffer [pH 7.4], 130 mM NaCl, 5 mM KCl, 1.4 mM CaCl₂, 1mM MgCl₂, 5.6 mM glucose, and 0.1% BSA). To measure FcεRI expression ex vivo, MCs were enriched from peritoneal lavage by 70% Percoll Gradient (GE Healthcare, UK) and stained with APC anti-CD45 Ab (clone 104), FITC anti-c-kit Ab (clone 2B8) and PE anti- FcεRI Ab (clone MAR-1), all from eBioscience.

β-hexosaminidase and cytokine release assay and PLC-γ2 phosphorilation

IgE pre-sensitized BMMCs were challenged in Tyrode’s buffer with Ag (100ng/ml) for 30 min in the presence or absence of equal amount of the indicated cell types. In some experiments BMMCs and T cells were separated by a 3.0 μm Transwell membrane (Corning Life Sciences, Acton, MA). In some experiments pre-sensitized BMMCs were incubated for 30 min with 10μg/ml blocking anti-OX40L (clone MGP34) (Murata et al., 2000) or isotype control (rat IgG2c) Ab before Ag challenge in presence of CD4⁺CD25⁺ T_regs. Samples were placed on ice and immediately centrifuged to pellet cells. The enzymatic activities of β-hexosaminidase in supernatants and in the cell pellets, after solubilizing with 0.5% Triton X-100 in Tyrode’s buffer, were measured with p-nitrophenyl N-acetyl-β-D-glucosaminide in 0.1 M sodium citrate (pH 4.5) for 60 min at 37°C. The reaction was stopped by addition of 0.2 M glycine (pH 10.7). The release of the product 4-p-nitrophenol was detected by absorbance at 405 nm. The extent of degranulation was calculated as the percentage of 4-p-nitrophenol absorbance in the supernatants over the sum of absorbance in the supernatants and in cell pellets solubilized in detergent. For cytokine analysis, IgE-sensitized BMMCs and Tregs were cultured alone or together for 16 h in presence of 100ng/ml Ag. Concentrations of TNF-α and IL-6 were determined in supernatants using Mouse Inflammation Kit (BD Biosciences, San Diego, CA). To assess PLC-γ2 phosphorilation, BMMCs were Ag-stimulated in presence of T_reg and fixed after the indicated time, immediately stained with a PE-conjugated anti-PLC-γ2 (pY759) Ab (BD Bioscience, San Diego, CA) and FITC anti-c-kit Ab (clone 2B8, eBioscience). Flow cytometry data were acquired on a FACSCalibur (Becton Dickinson) and analyzed with FlowJo software (version 8.5.2; Treestar Inc., Ashland, OR).

Expression of mouse OX40 by K562 cells and preparation of membranes

Human chronic myelogenous leukemia K562 cells were stably transfected with empty pCDNA3 plasmid or pCDNA3 expressing murine OX40 molecule (OX40Dir 5′GCGAATTCAGAAAGCAGACAAGG3′; OX40Rev 5′CACTCGAGTACTAATGCTCAGAT 3′). OX40 positive K562 cell clones were identify by flow cytometry with anti-mouse OX40 antibody (OX86, BD Biosciences). Membranes from K562 and K562-OX40 cell clones were prepared as previously described (Merluzzi et al., 2008) and resuspended at a final concentration of 5 × 10⁷ cell equivalents per ml based on the starting cell numbers. Membranes were added to IgE-sensitized BMMCs at dilution of 1:125v/v together with the Ag.

Intracellular Ca²⁺ determination

For Ca²⁺ measurements, 1 × 10⁶ IgE-sensitized BMMCs were loaded with 3μM FURA-2AM (Molecular Probes, Eugene, OR) in RPMI 2% FBS for 45 min at 37°C. Cells were washed in Tyrodes-BSA buffer (Saitoh et al., 2000), incubated in the presence of equal amounts of the indicated cell types and challenged with Ag (20 ng/ml). All fluorescence measurements (excitation and emission wavelengths, 340/380 and 505 nm, respectively) were performed in a Perkin-Elmer LS-50B spectrofluorimeter (Perkin-Elmer, Norwalk, CT) equipped with a thermostatically controlled cuvette holder and magnetic stirring. During the experiment, temperature was kept at 37°C. The changes in [Ca2⁺]i are expressed as a ratio of the light emitted at 505 nm upon excitation at the two wavelengths, 340 and 380 nm (F340/F380).

FACS-based cell sort and cAMP ELISA

To evaluate the intracellular levels of cAMP, pre-sensitized BMMCs were labeled for 15 min with 5μM carboxyfluorescein diacetate, succinimidyl ester (CFDA-SE; CFSE) (Molecular Probes, Invitrogen, UK) and Ag-challenged in presence of T_regs. After co-culture, CFSE-labeled BMMCs and CD25PE-labeled T_regs were isolated using a MoFlo cell sorter (Dako). The purity of the sorted populations was >99%. As positive control, sensitized BMMCs were incubated with 25 μM forskolin for 1h before Ag stimulation. Cells were washed twice in PBS and lysed in 0,1M HCl/0,1% Triton X-100 (10⁷/ml). cAMP levels were measured using Correlate EIA Direct cAMP assay (Assay Design, Ann Arbor, MI, USA).

Systemic Anaphylaxis

Mice were sensitized with 3 μg of mouse DNP-specific IgE by tail vein injection. 24 h later, mice were challenged i.v. with 0.5 mg of Ag or vehicle (PBS). After 1.5 min, mice were euthanized with CO₂ and blood was withdrawn by cardiac puncture. Plasma histamine concentration was determined by ELISA according to the producer instruction (DRG Instruments GmbH, Germany).

This work was supported by grants from the Ministero dell’Istruzione Università e Ricerca (PRIN 2005), Agenzia Spaziale Italiana. (Progetto OSMA), AIRC (Associazione Italiana Ricerca sul Cancro) and LR.11 del Friuli Venezia Giulia. SP is supported by a fellowship from FIRC (Fondazione Italiana Ricerca sul Cancro). The work of JR and SO was supported by the National Insitute of Arthritis, Musculoskeletal and Skin Diseases of the National Institutes of Health. We thank Arlene H. Sharpe (Harvard) for providing OX40L^−/− bone marrow and Ivano Arioli for technical assistance. We also thank Daniela Cesselli and Elisa Puppato for cell sorting. The authors declare that they have no competing financial interests.