Zinc transporter Znt5/Slc30a5 is required for the mast cell–mediated delayed-type allergic reaction but not the immediate-type reaction

There are eight known members of the Znt family (45). To examine the role of the Znts in mast cells, we checked the expression level of each family member in BMMCs by examining the ReFDIC (Reference Database of Immune Cells) (46). The database showed that Znt5 was among the Znts that were highly expressed in BMMCs (Fig. S1). The messenger RNA (mRNA) level of Znt5 was also enhanced by FcεRI stimulation (Fig. 1 A and Fig. S2). Because Znt5^−/− mice are available (44), we focused on Znt5 to clarify its role in mast cell function. First, we determined that Znt5 protein was expressed in BMMCs from WT but not Znt5^−/− mice (Fig. 1 B). We also determined that in the BMMCs, Znt5 was mainly located in the Golgi (Fig. 1 C), as reported previously for other cell types (47, 48). Next, we investigated whether Zn homeostasis was disordered in Znt5^−/− mast cells using a cell-permeable Zn-specific fluorescent probe, Newport green DCF diacetate, which predominantly localizes to the cytoplasm. The Znt5^−/− BMMCs showed an increased level of free Zn in the cytoplasm (Znt5^+/+ = 0.0723 ± 0.00225 µM; Znt5^−/− = 0.106 ± 0.0132 µM; P < 0.05; Fig. 1 D), indicating that the intracellular distribution of free Zn was impaired in the Znt5^−/− BMMCs.

We next asked if Znt5 was required for mast cell development. Culturing BM cells from WT and Znt5^−/− mice in the presence of IL-3 yielded highly purified mast cell populations. These BMMCs, whether derived from WT or Znt5^−/− mice, were indistinguishable in morphology when stained with Alcian blue (Fig. S3 A). The growth rate and total cell numbers in these cultures, as well as the frequency of BMMCs as revealed by flow cytometric analysis of the surface expression of c-kit and FcεRI, were equivalent (Fig. S3 B). In addition to the normal in vitro development of the mast cells, the number of mast cells, their morphology, and their anatomical distribution in the ear, skin, and stomach of WT and Znt5^−/− mice were comparable (Fig. S4, A and B). Collectively, these findings demonstrated that Znt5 is not required for mast cell development.

We examined the roles played by Znt5 in vivo in PCA and CHS using the Znt5^−/− mice. We first tested the importance of Znt5 in the PCA model by evaluating the extravasation of Evans blue dye in the ears of mice that had been sensitized and challenged with an antigen. However, there was no notable difference in the extravasation kinetics or total amount of Evans blue dye in the ear in the WT and Znt5^−/− mice (Fig. 2 A), indicating that the mast cell–mediated PCA reaction occurs normally in the absence of Znt5.

We next examined the CHS response, which is promoted by mast cell–derived proinflammatory cytokines. WT mice developed robust CHS responses to experimental hapten such as FITC, as assessed by tissue swelling at the site of hapten challenge, which reached a maximum 24 h after antigen stimulation (Fig. 2 B). Consistent with a previous study (31), mast cell–deficient Kit^W-sh/W-sh mice developed a low-level FITC-induced CHS reaction (Fig. 2 B). The Znt5^−/− mice exhibited a similar defective CHS reaction (Fig. 2 B). Even 24 h after the antigen challenge, the increase in ear thickness of the Znt5^−/− mice was <45% of that of WT mice. Consistent with this result, the affected ear of the Znt5^−/− mice showed significantly reduced FITC-induced cytokine production and transcription of the genes for IL-6, IL-1β, MCP, and TNF-α (Fig. 2, C and D). The Znt5^−/− ears also showed a decrease in the transcription level of the genes for chemokines such as MIP-2 (CXCL2; Fig. 2 C). From these results, we concluded that Znt5 was selectively required for CHS, but not for PCA, in vivo. To determine whether defectiveness in CHS response in Znt5^−/− mice is caused by the lack of Znt5 in mast cell but not in other environmental cells and tissues, we checked Kit^W-sh/W-sh mice that had been engrafted with mast cell from either WT or Znt5^−/− mice. We found that engraftment with BMMCs from WT mice, but not Znt5^−/− mice, restored CHS response in Kit^W-sh/W-sh mice (Fig. 2 E). Consistent with these results, Kit^W-sh/W-sh mice reconstituted with Znt5^−/− BMMCs had significantly reduced FITC-induced cytokine expression level such as TNF-α and IL-6 in the ear (Fig. 2 F). The number of mast cells per mm² of dermis did not change between Znt5^+/+ and Znt5^−/− BMMC-engrafted Kit^W-sh/W-sh mice (Fig. S5). Collectively, these findings showed that Znt5 in mast cells is required for FITC-induced CHS responses.

FcεRI-induced degranulation and the production of lipid mediators play crucial roles in the PCA reaction, whereas cytokine production is essential for the CHS response (49). To learn the basis for the selective defect in the CHS reaction of Znt5^−/− mice, we examined the capacities of Znt5^−/− BMMCs to degranulate, synthesize, and secrete lipid mediators and to produce cytokines. BMMCs derived from Znt5^−/− mice released similar amounts of β-hexosaminidase to WT BMMCs in response to FcεRI stimulation (Fig. 3 A), indicating that their ability to degranulate was intact. Furthermore, FcεRI stimulation-induced lipid mediator secretion, such as that of leukotrienes C₄, occurred normally in the Znt5^−/− BMMCs (Fig. 3 B). These results were consistent with the normal PCA reaction observed in the Znt5^−/− mice. However, the production of the FcεRI-induced cytokines IL-6 and TNF-α was significantly reduced in Znt5^−/− BMMCs as compared with WT BMMCs (Fig. 3 C). Furthermore, PMA-induced cytokine production was also diminished in Znt5^−/− BMMCs (Fig. 3 D). To rule out the possibility that the defect in cytokine production was caused by an impaired cytokine transport step in Znt5^−/− BMMCs, we checked the ability of LPS to induce cytokine production. As shown in Fig. 3 E, the LPS-induced production of the cytokines IL-6 and TNF-α did not decrease in the Znt5^−/− BMMCs. These results indicated that Znt5 is selectively required for FcεRI-induced cytokine production but not for degranulation in BMMCs.

To define the molecular mechanisms responsible for the defective cytokine production in Znt5^−/− BMMCs, we analyzed the cytokine mRNA levels using quantitative PCR. The FcεRI stimulation of BMMCs led to a time-dependent increase in the IL-6 and TNF-α mRNA level. However, the level of IL-6 and TNF-α mRNA that was induced was significantly lower in the Znt5^−/− BMMCs (Fig. 4 A). As expected from the result shown in Fig. 3 E, the levels of LPS-induced IL-6 and TNF-α mRNA in the Znt5^−/− BMMCs were not decreased as compared with those in WT (Fig. 4 B). Thus, Znt5 contributed to the FcεRI-induced increase of mRNA for IL-6 and TNF-α.

We further analyzed whether Znt5 was involved in the FcεRI-induced nuclear translocation of NF-κB. As shown in Fig. 4 C, the FcεRI-induced translocation of p65 to the nucleus was significantly decreased in the Znt5^−/− BMMCs. Consistent with these results, FcεRI-mediated IκBα phosphorylation and its subsequent degradation were impaired in the Znt5^−/− BMMCs (Fig. 4 D and Fig. S6). In contrast, the FcεRI-induced activation of the mitogen-activated protein kinases (MAPKs) ERK1/2, JNK1/2, and p38 was not diminished in the Znt5^−/− BMMCs (Fig. 4 E). We also investigated whether Znt5 was involved in FcεRI-induced calcium-elicited signaling events. We monitored the tyrosine phosphorylation of LAT and PLC-γ1, which contributed calcium signaling (26, 50). As expected from the result shown in Fig. 3 A, LAT and PLC-γ1 tyrosine phosphorylation was apparently normal in Znt5^−/− BMMCs, as was the FcεRI-induced calcium influx (Fig. S7, A and B). Collectively, these results show that Znt5 was selectively required for FcεRI-mediated NF-κB activation in BMMCs.

Because IκBα phosphorylation decreased in Znt5^−/− BMMCs (Fig. 4 D and Fig. S6), we next sought to identify the upstream target of Znt5 that was involved in NF-κB activation. Because FcεRI-induced NF-κB activation is dependent on PKC activation (23, 51), we examined the effect of Znt5 on the translocation of PKC to the plasma membrane, a process important for PKC function. As shown in Fig. 5 (A and B), the FcεRI-induced plasma membrane translocation of PKC-β was diminished in the Znt5^−/− BMMCs, as was the PMA-induced plasma membrane translocation of PKC-β, and a similar result was obtained in the case of plasma membrane translocation of PKC-α (not depicted). In addition, activity of PKC was reduced in the Znt5^−/− BMMCs compared with that in Znt5^+/+ BMMCs (Fig. S8). Furthermore, we confirmed that transfected Znt5 complementary DNA (cDNA) into the Znt5^−/− BMMCs rescued the defect in FcεRI-induced plasma membrane translocation of PKC-β (Fig. 5 C). To rule out the possibility that Znt5 altered the PKC-β expression level, we checked the protein level after FcεRI stimulation and did not find any decrease compared with the level in WT BMMCs (Fig. S9). Together, these results showed that Znt5 is required for the FcεRI-mediated translocation of PKC-β to the plasma membrane and that this effect on PKC-β translocation is, at least in part, the mechanism underlying involvement of Znt5 in FcεRI-induced NF-κB activation, which results in cytokine production.

To understand the mechanisms by which Znt5 is involved in the plasma membrane translocation of PKC, we focused on the C1 domain of PKC, the DAG-binding site. The C1 domain of PKC has two Zn finger–like motifs (Fig. 6 A) (39, 40). Based on the known function of the Zn finger, we hypothesized that the Zn finger–like motifs of PKC are required for translocation of PKC-β to the plasma membrane. To demonstrate this hypothesis, we designed WT–PKC-β–GFP and three mutant PKC-β–GFP constructs (Fig. 6 A) and examined their ability to undergo PMA-induced plasma membrane translocation in HeLa cells.

The WT and C1BS mutant form of PKC-β–GFP translocated equally well to the plasma membrane upon stimulation by PMA (Fig. 6 B and Videos 1 and 3). In contrast, the C1AS and C1ABS double mutant constructs did not translocate well and were retained in the cytoplasm (Fig. 6 B and Videos 2 and 4). These data suggested that the C1A Zn finger–like motif of PKC is important for its translocation to the plasma membrane and that this motif is a possible target for Znt5 and the likely binding site for DAG.

To test this hypothesis, we designed a biotin-conjugated PMA, an analogue of DAG (Biotinyl-PMA) (Fig. S10 A). Biotinyl-PMA induced cytokine production, just as PMA did (Fig. S10 B). Using Biotinyl-PMA, we assessed the binding ability of PMA to PKC-β. Transfected PKC-β WT and the C1AS mutant were extracted, run on a gel, and electrotransferred to a polyvinylidine fluoride (PVDF) membrane under native conditions. As shown in Fig. 6 C, Biotinyl-PMA expected to bind the PKC WT construct, but its signal intensity to the PKC C1AS mutant form was lower. Consistent with this result, the signal intensity of Biotinyl-PMA to PKC obtained from Znt5^−/− BMMCs was less than that to PKC from WT BMMCs (Fig. 6 D).

In this paper, we demonstrate that a Znt, Znt5, plays a crucial role in mast cell activation and mast cell–mediated allergic reactions. The mast cell is widely recognized as the most important effector cell in IgE-associated allergic disorders such as anaphylaxis. Previous experiments with genetically mast cell–deficient Kit^W/Kit^W-V or Kit^W-sh/Kit^W-sh mice have shown that mast cells are required for the IgE-mediated immediate-type allergic response (24, 27). Thus, the participation of mast cells in the immediate allergic reaction is well established. In addition to this classical role, much evidence has accumulated showing that mast cells play crucial roles in delayed-type allergic reactions and chronic inflammatory diseases, including arthritis, experimental allergic encephalomyelitis, colitis, and CHS (28, 52–54). A variety of mast cell–derived cytokines and chemokines are likely to be essential for delayed-type allergic reactions and chronic inflammatory diseases. In fact, the mast cell–defective Kit^W/Kit^W-V mouse does not develop experimental allergic encephalomyelitis, and this defect is rescued by transferred WT BMMCs but not by IL-4–deficient BMMCs (55). These studies indicate that mast cell–derived cytokines are important in inflammatory diseases.

In this paper, we showed that Znt5^−/− mast cells had an intact degranulation process but impaired cytokine and chemokine production. Most importantly, Znt5^−/− mice had a defective mast cell–dependent delayed-type allergic reaction, i.e., CHS, but their immediate-type reaction, i.e., PCA, was unimpaired. Therefore, the Znt5^−/− mouse will be valuable for further dissection of the in vivo roles of mast cells in immune and allergic responses and for analyzing the pathological status of a variety of immune-related diseases.

Our results showed that Znt5 is important for FcεRI-mediated cytokine or chemokine production. As Znt5 was mainly located in the Golgi, we first examined whether Znt5^−/− mast cells were defective in cytokine processing/secretion. However, LPS-induced cytokine production was normal, indicating that the transport step was unaffected by the Znt5 deficiency in mast cells. We also observed that FcεRI induced normal activation of LAT and PLC-γ and calcium mobilization. This observation is consistent with our result showing normal degranulation in Znt5^−/− BMMCs. From this, we deduced that the degranulation process can be induced in Znt5^−/− mast cells as a result of intact calcium influx. Our results do not dispute the reported involvement of PKC in the degranulation process (35, 56), but this requirement for PKC may be compensated by other signaling pathways in Znt5^−/− mast cells. Alternatively, the threshold of the degree of PKC activation required for degranulation and that for NF-κB activation might be different. To support this idea, we performed PKC family inhibitor experiment and showed that PKC inhibitor-treated Znt5^−/− mast cells were significantly reduced by FcεRI-mediated degranulation (Fig. S11).

We found that FcεRI-mediated NF-κB activation was defective in the Znt5^−/− BMMCs; FcεRI-induced nuclear translocation of p65 was reduced, and FcεRI-mediated IκBα phosphorylation and its subsequent degradation were impaired. From these results, we concluded that Znt5 is selectively required for FcεRI-mediated NF-κB activation but not for MAPK activation or calcium mobilization. Because the expression of IL-6 and TNF-α depend on the activation of NF-κB, which is a master transcription factor that regulates the expression of proinflammatory cytokine and chemokine genes (57), our results indicated that Znt5 is a novel player involved in FcεRI-induced cytokine production through NF-κB activation.

PKC-θ and PKC-β play important roles in NF-κB activation and function upstream of the Bcl10–MALT1 complex in TCR and BCR signaling (41, 42). In mast cells, PKC is responsible for the activation of NF-κB; for example, a PKC inhibitor suppresses NF-κB activation (23, 51). Furthermore, PKC-β–deficient mast cells are defective in IL-6 production (58). In this paper, we showed that Znt5 was required for the FcεRI- or PMA-induced translocation of PKC-β to the plasma membrane of BMMCs, and this requirement explains why Znt5 is needed for NF-κB activation.

A related question is how Znt5 regulates the plasma membrane translocation of PKC-β, and several of our findings relate to this issue: the C1A Zn finger–like motif of PKC-β was required for translocation of PKC to the plasma membrane; mutating the C1A Zn finger–like motif of PKC-β diminished its ability to bind PMA; PKC obtained from Znt5^−/− BMMCs showed decreased binding to PMA; Znt5 was located in the Golgi; and Znt5^−/− BMMCs showed an increase in cytoplasmic free Zn. Because Znt5 was located in the Golgi, it probably promotes a Zn efflux from the cytoplasm into the Golgi. Because Znt5^−/− BMMCs had increased cytoplasmic free Zn, it seems likely that the available Zn in the Golgi was decreased concomitantly.

Together, these observations suggested the hypothesis that in the Znt5^−/− BMMCs, the Zn available to the C1A Zn finger–like motif of PKC as it transits through the Golgi is decreased, which results in an increase in Zn-free PKC. The mutation of the Zn-binding site of C1A interfered with the plasma membrane translocation of PKC, and this mutation also diminished PMA binding to PKC. Collectively, our findings suggest that binding of Zn to the C1A of PKC induces a conformational change required for DAG binding, which is itself required for the translocation of PKC to the plasma membrane. This hypothesis is supported by our data as well as by findings from other groups that Zn is involved in the translocation of PKC to the plasma membrane (59–61). Furthermore, in experiments using Znt5^−/− DT40 cells, Suzuki et al. (48, 62) showed that Znt5 expressed on the ER-Golgi membrane is required for the enzymatic activity of the Zn-dependent alkaline phosphatases (ALPs), which are processed from apoALPs to holoALPs in the ER-Golgi.

In summary, the results in this paper show that a Znt, Znt5, is selectively required for the mast cell–mediated delayed-type allergic response but not for the immediate-type allergic reaction. We further showed that Znt5 is required in the PKC-mediated NF-κB signaling pathway. The identification of Znt5 as a novel component of FcεRI-mediated PKC activation sheds light on the molecular mechanisms of the processing and activation of PKC and of the Zn-mediated regulation of mast cell–dependent allergic responses, specifically of the delayed type.

C57BL/6J mice were obtained from CREA Japan. Znt5^−/− mice were provided by Y. Nakamura and T. Tanaka (University of Tokyo, Tokyo, Japan, and RIKEN, Center for Genomic Medicine, Kanagawa, Japan). The generation of Znt5^−/− mouse was reported in detail by Inoue et al. (44). Mast cell–deficient Kit^W-sh/W-sh mice were provided by the RIKEN BioResource Center. We obtained approval from the Animal Research Committee at RIKEN for all animal experiments performed.

Anti-Znt5 antibodies were raised by immunizing rabbits with KLH-conjugated peptide (TIQVEKEAYFQHMSG). For the immunization, rabbits were first injected with 200 µg of KLH-conjugated Znt5 peptide in complete Freund’s adjuvant and then boosted every week with 50 µg of the antigen in incomplete Freund’s adjuvant. Anti-GFP and antiphosphotyrosine (4G10) antibodies were purchased from MBL and Millipore, respectively. Anti-Flag (M2), anti-GM130, anti–PKC-β (C-16), anti–phospho-PKC (S660), and anti-LAT antibodies were obtained from Sigma-Aldrich, BD, Santa Cruz Biotechnology, Inc., Cell Signaling Technology, and Millipore, respectively. Gö6976 was purchased from EMD.

The FITC-induced CHS procedure was performed as described previously (31). In brief, mice were sensitized with a total of 200 µl of 2% FITC isomer-I (FITC; Sigma-Aldrich) in a vehicle consisting of acetone-dibutylphthalate (1:1) applied to the skin of the back. 5 d after the sensitization with FITC, the mice were challenged with 40 µl of vehicle alone on the right ear (20 µl on each side of the ear) and 0.5% FITC on the left ear (20 µl on each side). Ear thickness was measured before and at multiple intervals after FITC challenge with an engineer’s microcaliper (Ozaki). Some mice from each group were killed 24 h after the FITC or vehicle challenge for cytokine analysis.

For mast cell reconstitution studies, BMMCs were injected intradermally (1.0 × 10⁶ cells in 20 µl/ear for CHS) in 6-wk-old Kit^W-sh/W-sh mice. The mice were used for CHS experiments at 6 wk after transfer of BMMCs.

A total of 2 µg IgE was injected subcutaneously into ears for 12 h. After this sensitization, the mice were then challenged with an intravenous injection of 250 µg of polyvalent DNP-BSA (Cosmo Bio Co., LTD.) in 250 µl of saline and 5 mg/ml of Evans blue dye (Sigma-Aldrich). The extravasation of Evans blue into the ear was monitored for 30 min. The mice were then sacrificed, both ears were dissected, and the Evans blue dye was extracted in 700 µl of formamide at 63°C overnight. The absorbance of the Evans blue–containing formamide was then measured at 620 nm. To observe the ear mast cells, 3-µm paraffin sections were fixed and stained with nuclear fast red and Alcian blue, and the Alcian blue–stained cells in each sample were counted. All data are expressed as the number of mast cells per mm² of dermis.

BMMCs were prepared as described previously (63). In brief, BM cells were flushed from the marrow cavity of mouse femurs, and the mast cells (BMMCs) were selectively grown in RPMI1640 medium supplemented with IL-3 (conditioned medium from an mIL-3–producing cell line, CHOmIL-3-3-12M [gift from T. Sudo, Toray Industries, Inc., Kamakura, Japan]), 10% FBS, 10 mU/ml penicillin, and 0.1 mg/ml streptomycin for 4–8 wk. During culture, the medium was changed every 3–4 d, and the cells were transferred to new dishes to eliminate adherent cells. After 5 wk in culture, the BMMCs were ready for in vitro experiments and showed the cell surface expression of FcεRI and c-Kit. HeLa cells were maintained in RPMI1640 supplemented with 10% FBS, penicillin, and streptomycin.

The degranulation assay was described previously (64). In brief, the degree of degranulation was determined by measuring the release of β-hexosaminidase. 1 × 10⁶ cells/ml were preloaded with 1 µg/ml anti-DNP IgE (SPE-7; Sigma-Aldrich) by a 6-h incubation in medium (without IL-3). To measure β-hexosaminidase release, the sensitized cells were stimulated with 100 ng/ml DNP-HSA for 30 min in Tyrode’s buffer. Samples were placed on ice and then spun at 4°C for 5 min. The enzymatic activities of β-hexosaminidase in the supernatants and cell pellets solubilized with 1% Triton X-100 in Tyrode’s buffer were measured with p-nitrophenyl N-acetyl-β-d-glucosaminide (Sigma-Aldrich) in 0.1 M sodium citrate, pH 4.5, for 60 min at 37°C. The reaction was stopped by the 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 by dividing the 4-p-nitrophenol absorbance in the supernatant by the sum of the absorbance in the supernatant and detergent-solubilized cell pellet.

Cells were activated as described in the previous sections, and TNF-α and IL-6 in the cell culture supernatants were measured with an ELISA kit (eBioscience). The leukotrienes were quantified using an ELISA kit (Cayman Chemical), according to the manufacturer’s instructions.

1 × 10⁶/ml BMMCs were sensitized with 1 µg/ml IgE for 6 h at 37°C. Cells were stimulated with 100 ng/ml DNP-HSA. After the stimulation, the cells were harvested and lysed with 100 µl of lysis buffer (20 mM Tris-HCl, pH 7.4, at 4°C, 150 mM NaCl, 1% NP-40, 0.1% SDS, 0.1% deoxycholate, 1 mM NaVO₄, 3 mM EDTA, and proteinase inhibitors [0.5 mM PMSF, 10 µg/ml aprotinin, 5 µg/ml pepstatin, and10 µg/ml leupeptin]) for 30 min at 4°C and spun at 15,000 g at 4°C for 15 min. The eluted and reduced samples were resolved by SDS-PAGE using a 5–20% gradient polyacrylamide gel (Wako Chemicals USA, Inc.) and transferred to a PVDF membrane (Immobilon-P; Millipore). For immunoblotting, the membranes were incubated with anti–phospho-ERK, anti–phospho-JNK, anti–phospho-p38, anti-ERK, anti-JNK, anti-p38, anti–phospho-IκBα, anti-IκBα, anti–β-tubulin, anti-actin, and anti–PKC-β, respectively. For immunoprecipitation, the cell lysates were incubated for 4 h at 4°C with 1 µg of antibodies bound to protein A Sepharose. The eluted and reduced samples were separated on a 5–20% gradient SDS-polyacrylamide gel and transferred to a PVDF membrane. The membranes were incubated with antiphosphotyrosine. After blotting with the first antibody, the membranes were blotted with HRP-conjugated species-specific anti-IgG (Invitrogen) for 1 h at room temperature. After extensive washing of the membranes, the immunoreactive proteins were visualized using the Renaissance chemiluminescence system (Dupont), according to the manufacturer’s recommendations. The PVDF membranes were exposed to RX film (Fujifilm). Densitometric analysis for tubulin and PKC-β was performed using a fluorescence image analyzer (LAS-1000; Fujifilm).

Plasma membrane proteins were extracted using a Plasma Membrane Protein Extraction kit (K268-50; BioVision, Inc.), according to the manufacturer’s instructions. The plasma membrane fractions were used for immunoblotting and probed with anti-actin, anti–PKC-β, and anti–phospho-PKC (S660). Densitometric analysis for PKC-β, phospho-PKC (S660), and actin was performed using an LAS-1000 fluorescence image analyzer.

Separate aliquots of 5 × 10⁵ cells were sensitized with 1 µg/ml IgE for 6 h. The cells were then stimulated with 100 ng/ml DNP-HSA for 10 min at 37°C and fixed with 4% paraformaldehyde for 30 min. The cells were spun and permeabilized in Perm Buffer (BD) containing 1% BSA for 15 min at room temperature. The cells were washed with 1 ml PBS⁻ (PBS without calcium and magnesium) twice, resuspended in 500 µl of PBS⁻, and attached to glass slides by cytospin (Thermo Fisher Scientific) for 5 min. Primary and secondary staining were performed on the slides, with anti-p65 at a dilution of 1:50, anti–PKC-β at 1:100, Alexa Fluor 488–conjugated anti–rabbit IgG (Invitrogen) at 1:100, and phalloidin-rhodamine (Invitrogen) at 1:100. Confocal microscopy was performed using the TCS SL system (Leica).

Cells were homogenized with Sepasol RNAI (Nacalai Tesque, Inc.), and the total RNA was isolated according to the manufacturer’s instructions. For standard RT-PCR, cDNA was synthesized from 500 ng of total RNA by RT (ReverTra Ace; TOYOBO) and 500 ng of oligo (dT) primer (Invitrogen) for 30 min at 42°C. A portion of the cDNA (typically 1/20 vol) was used for standard PCR to detect IL-6, TNF-α, and GAPDH. 25 cycles of PCR were performed with 0.5 U rTaq DNA polymerase and 10 pmol of gene-specific sense and antisense primers. For real-time quantitative RT-PCR, IL-6 and TNF-α gene expression was measured relative to GAPDH using the SYBR Green system (Applied Biosystems). Primers used in these experiments were purchased from Invitrogen, and the sequences were as follows: IL-6 forward, 5′-GAGGATACCACTCCCAACAGACC-3′ and reverse, 5′-AAGTGCATCATCGTTGTTCATACA-3′; TNF-α forward, 5′-CATCTTCTCAAAATTCGAGTGACAA-3′ and reverse, 5′-TGGGAGTAGACAAGGTACAACCC-3′; GAPDH forward, 5′-TTCACCACCATGGAGAAGGCCG-3′ and reverse, 5′-GGCATGGACTGTGGTCATGA-3′; and hypoxanthine-guanine phosphoribosyl transferase forward, 5′-CCTCCCATCTCCTTCATGACA-3′ and reverse, 5′-GATTAGCGATGATGAACCAGGTT-3′. In addition, primers used for amplifying cDNAs for Znt family members and other primer sets are listed in Table S1. PCR products were separated on an agarose gel, stained with 500 ng/ml ethidium bromide, and photographed.

2 × 10⁶ BMMCs were treated with 2.5 µM of Newport green for 30 min at 37°C. The cells were washed two times with PBS⁻ before the fluorescence intensity was measured by FACS. The concentration of intracellular free Zn was calculated from the mean fluorescence with the formula [Zn] = K_D × [(F − F_min)/(F_max − F)]. The dissociation constant of the Newport Green/Zn complex is 1 µM. F_min was determined by the addition of the Zn-specific membrane-permeant chelator TPEN, and F_max was determined by the addition of ZnSO₄ and the ionophore pyrithione (65).

1 × 10⁶ BMMCs/ml were sensitized with 1 µg/ml IgE (anti-DNP IgE clone SPE-7; Sigma-Aldrich) for 6 h at 37°C. IgE-sensitized mast cells were washed three times, resuspended in Tyrode’s buffer (10 mM Hepes, pH 7.4, 130 mM NaCl, 5 mM KCl, 1.4 mM CaCl₂, 1 mM MgCl₂, and 5.6 mM glucose), allowed to adhere to a poly-l-lysine–coated glass-bottom dish, and incubated with 5 µM Fluo-4, a cell-permeant dye, for 30 min at 37°C. Surplus fluorescence indicator and floating cells were removed by at least three washes with Tyrode’s buffer. Cells were stimulated with 100 ng/ml DNP–HSA (Sigma-Aldrich) at 37°C. Images of the fluorescent signals were captured every 10 or 30 s from an inverted microscope (Axiovert 200 MOT; Carl Zeiss, Inc.) equipped with an oil Plan Neofluar 100× 1.3 NA objective, a charge-coupled device camera (Cool Snap HQ; Roper Scientific), and the system control application SlideBook (Intelligent Imaging Innovation) at 25°C. The captured images were processed with Photoshop software (Adobe) to adjust their size and contrast.

PKC-β was cloned by PCR using cDNA from BMMCs and sequenced on a 3130xl sequencer (ABI-PRISM; APplied Biosystems). The GFP-tagged expression vectors for PKC-β were constructed and subcloned into pEGFP N1 (Invitrogen). The construct encoding mutant PKC-β–GFP, in which the Zn finger–like motif in the C1 region was mutated, was made using a PCR-based method. The human Znt5 cDNA in pA-Zeocin was reported in detail by Suzuki et al. (62). HeLa cells were transfected with a total of 1 µg PKC-β cDNA in pEGFP or with empty vector using Lipofectamine 2000 (Invitrogen).

Retroviral infection was performed as previously described (64). The Flag-tagged human Znt5 plasmid (gift from T. Kambe and T. Suzuki, Kyoto University, Kyoto, Japan) was inserted into the BamHI and NotI sites of the retroviral vector pMX (gift from T. Kitamura, University of Tokyo, Tokyo, Japan). This construct was then used to transfect the 293T-based packaging cell line phoenix (gift from G. Nolan, Stanford University, Stanford, CA), with Lipofectamine 2000 (Invitrogen) to generate recombinant retroviruses. BM cells were infected with the retrovirus in the presence of 10 µg/ml polybrene (Sigma-Aldrich) and IL-3.

PMA (Nacalai Tesque, Inc.) was conjugated with biotin (Wako Chemicals USA, Inc.). The predicted biotinyl-PMA is shown in Fig. S10. Native PAGE was performed with the NativePAGE Novex Bis-Tris Gel system (Invitrogen). In detail, the cells were lysed with NativePAGE Sample buffer for 10 min at 4°C and spun at 15,000 g at 4°C for 10 min. The samples were separated by native PAGE on 4–16% Bis-Tris Gels and transferred to a PVDF membrane (Immobilon-P; Millipore). For immunoblotting, the membrane was incubated with Biotinyl-PMA or anti–PKC-β antibody. After the membrane was washed in TBST, the coloring reaction was performed with streptavidin-AP (Roche) or anti–rabbit IgG HRP antibody, respectively. Densitometric analysis for PKC-β and PMA was performed using an LAS-1000 fluorescence image analyzer (Fujifilm).

All data were analyzed with Statcell (Microsoft). Data were considered statistically significant for p-values <0.05, obtained with a two-tailed Student’s t test.

Fig. S1 shows expression level of Znt family member in BMMCs. Fig. S2 shows expression level of Znt5 after antigen stimulation. Fig. S3 shows normal development of BMMC from Znt5^−/− mice. Fig. S4 shows normal development of mast cell in Znt5^−/− mice. Fig. S5 shows numbers of mast cells in dermis from vehicle-treated or FITC-challenged ears. Fig. S6 shows quantitative analysis of IκBα phosphorylation and degradation. Fig. S7 shows normal FcεRI-induced calcium signaling and influx in Znt5^−/− BMMCs. Fig. S8 shows impairment of PKC activity in Znt5^−/− BMMCs. Fig. S9 shows normal expression levels of PKC-β in Znt5^−/− BMMCs. Fig. S10 shows design of biotinyl-PMA and that biotinyl-PMA can induce cytokine production. Fig. S11 shows that the PKC inhibitor Gö6976 inhibits FcεRI-induced degranulation in Znt5^−/− BMMCs. Video 1 shows time-lapse images of WT PKC-β–EGFP in HeLa cells after PMA stimulation. Video 2 shows time-lapse images of C1AS PKC-β–EGFP in HeLa cells after PMA stimulation. Video 3 shows time-lapse images of C1BS PKC-β–EGFP in HeLa cells after PMA stimulation. Video 4 shows time-lapse images of C1ABS PKC-β–EGFP in HeLa cells after PMA stimulation.

We thank Drs. Y. Nakamura and T. Tanaka (University of Tokyo and RIKEN, Center for Genomic Medicine) for providing the Znt5^−/− mice. We would like to thank Drs. P. Burrows for critical readings and K. Kabu, T. Suzuki, T. Kambe, A. Hijikata, H. Shinohara, H. Kitamura, and H. Watarai for technical advice and suggestions. We also thank Ms. A. Ito, Ms. M. Yamada, Ms. M. Hara, Mr. S. Fukuda, and Ms. N. Hashimoto for excellent technical assistance and Ms. M. Shimura for secretarial assistance.

This work was supported by grants from the Ministry of Education, Culture, Sports, Science and Technology in Japan.

The authors have no conflicting financial interests.