Ca2+-mediated signal transduction pathways play a central regulatory role in dendritic cell (DC) responses to diverse Ags. However, the mechanisms leading to increased [Ca2+]i upon DC activation remained ill-defined. In the present study, LPS treatment (100 ng/ml) of mouse DCs resulted in a rapid increase in [Ca2+]i, which was due to Ca2+ release from intracellular stores and influx of extracellular Ca2+ across the cell membrane. In whole-cell voltage-clamp experiments, LPS-induced currents exhibited properties similar to the currents through the Ca2+ release-activated Ca2+ channels (CRAC). These currents were highly selective for Ca2+, exhibited a prominent inward rectification of the current-voltage relationship, and showed an anomalous mole fraction and a fast Ca2+-dependent inactivation. In addition, the LPS-induced increase of [Ca2+]i was sensitive to margatoxin and ICAGEN-4, both inhibitors of voltage-gated K+ (Kv) channels Kv1.3 and Kv1.5, respectively. MHC class II expression, CCL21-dependent migration, and TNF-α and IL-6 production decreased, whereas phagocytic capacity increased in LPS-stimulated DCs in the presence of both Kv channel inhibitors as well as the ICRAC inhibitor SKF-96365. Taken together, our results demonstrate that Ca2+ influx in LPS-stimulated DCs occurs via Ca2+ release-activated Ca2+ channels, is sensitive to Kv channel activity, and is in turn critically important for DC maturation and functions.
Dendritic cells (DCs)4 are essential for initiating and directing Ag-specific T cell responses. Ca2+-mediated signal transduction pathways play a critical regulatory role in DC responses to diverse Ags, including TLR ligands, intact bacteria, and microbial toxins (1). The phospholipase C-Ca2+ pathway seems to be involved in the maturation of monocyte-derived DCs induced by different agonists such as LPS, cholera toxin, dibutyryl-cAMP, and PGE2 as suggested from the effects of intracellular and extracellular Ca2+ chelation (2). Furthermore, addition of Ca2+ ionophores to immature DCs results in the acquisition of many morphological and functional properties of activated mature DCs (2, 3, 4).
Changes in the cytosolic Ca2+ concentration [Ca2+]i regulate receptor-mediated endocytosis, phagosome-lysosome fusion, and Ag processing. Upon DC stimulation by diverse soluble and particulate Ags, Calmodulin kinase (CamK) II is activated and inhibition of CamK II results in suppression of cytokine production (5). Another Ca2+-dependent kinase, CamK IV, which is also expressed in DCs, plays a key role in the pathway linking TLR-4 to the control of DC life span (6).
In a number of studies, stimulus-induced changes in [Ca2+]i have been observed. Thus, stimulation of DCs by lysophosphatidic acid resulted in a rapid increase in [Ca2+]i, which was not dependent on the presence of extracellular Ca2+ (7). Similarly, ligation of DC-SIGN, a C-type lectin in DCs that mediates capture and internalization of viral, bacterial, and fungal pathogens, triggered rapid and transient intracellular Ca2+ mobilization (8). However, the mechanisms and the consequences of this [Ca2+]i increase remain unclear.
In macrophages and Kupffer cells, LPS treatment causes an increase in [Ca2+]i which is related to TNF-α production (9, 10). Kupffer cells contain l-type voltage-dependent Ca2+ channels (11) and thus depolarization of the cell membrane is required for the Ca2+ influx into these cells. Contrary to Kupffer cells, DCs have been shown to possess Ca2+ release-activated Ca2+ channels (CRAC) as a main Ca2+ entry pathway (12). Accordingly, Ca2+ influx is enhanced by membrane hyperpolarization (12).
The present study explores the mechanism of Ca2+ entry into DCs. It is shown that similar to macrophages and Kupffer cells, DCs respond to LPS exposure with a transient increase in [Ca2+]i. We present evidence for the activation of ICRAC and demonstrate the importance of this activation for DC functions.
In addition, DCs were shown to contain voltage-gated K+ (Kv) channels, belonging to Kv1 channel family, which are up-regulated by LPS stimulation and play a role in cytokine production (13, 14). In the present study, we address the question whether Kv channels can modulate Ca2+ entry in DCs by maintaining the electrochemical driving force for Ca2+ influx (15) and thus participate in Ca2+-dependent DC functions.
Materials and Methods
DCs were cultured from bone marrow as previously described (16), with slight modifications. In brief, bone marrow cells were flushed out of the cavities from the femur and tibia of 7–11-wk-old female NMRI mice (Charles River Laboratories) with PBS. Cells were then washed twice with RPMI 1640 and seeded out at density of 2 × 106 cells per 60-mm dish. Cells were cultured for 6 days in RPMI 1640 (Life Technologies) containing: 10% FCS, 1% penicillin/streptomycin, 1% glutamine, 1% nonessential amino acids, and 0.05% 2-ME. Cultures were supplemented with GM-CSF (35 ng/ml, PeproTech) and fed with fresh medium containing GM-CSF on days 3 and 6. Nonadherent and loosely adherent cells were harvested after 6 days of culture. At day 7, >95% of the cells expressed CD11c, which is a marker for mouse DCs. Experiments were performed on mature DCs at days 7–9.
Immunostaining and flow cytometry
Cells (4 × 105) were incubated in 100 μl FACS buffer (PBS plus 0.1% FCS) containing fluorochrome-conjugated Abs at a concentration of 10 μg/ml. A total of 2 × 104 cells were analyzed. The following Abs (all from BD Pharmingen) were used for staining: FITC-conjugated anti-mouse CD11c, clone HL3 (Armenian Hamster IgG1, λ2), PE-conjugated anti-mouse CD86, clone GL1 (Rat IgG2a, κ), PE-conjugated rat anti-mouse I-A/I-E, clone M5/114.15.2 (IgG2b, κ) and PE-conjugated anti-mouse ICAM-1 (CD-54), clone 3E2 (Armenian Hamster IgG1, κ). After incubating with the Abs for 60 min at 4°C, the cells were washed twice and resuspended in FACS buffer for flow cytometry analysis.
Measurement of intracellular Ca2+
Fluorescence measurements were conducted with an inverted phase-contrast microscope (Axiovert 100, Zeiss). Cells were excited alternatively at 340 or 380 nm and the light was deflected by a dichroic mirror into either the objective (Fluar 40 × 1.30 oil, Zeiss) or a camera (Proxitronic). Emitted fluorescence intensity was recorded at 505 nm and data acquisition was accomplished by using specialized computer software (Metafluor, Universal Imaging). As a measure for the increase of cytosolic Ca2+ activity, the slope and peak of the changes in the 340/380 nm ratio were calculated for each experiment.
DCs were pretreated at 37°C with either the ICRAC inhibitor SKF-96365 (10 μM, 30 min; Sigma-Aldrich), or both margatoxin (MgTx, 1 nM, 30 min; Alomone Laboratories) and ICAGEN-4 (10 μM, 30 min; Ref. 17), inhibitors of Kv1.3 and Kv1.5, respectively, or left untreated. The cells were loaded with fura-2/AM (2 μM, Molecular Probes) for 30 min at 37°C. Intracellular Ca2+ was measured before and following addition of LPS from Escherichia coli (0.1 μg/ml; Sigma-Aldrich) in the absence or presence of extracellular Ca2+. When the cells were pretreated with channel inhibitors, the respective inhibitors were added to the bath solutions.
Alternatively, changes in cytosolic Ca2+ were monitored upon depletion of the intracellular Ca2+ stores. Experiments were conducted before and during exposure to Ca2+-free solution. In the absence of Ca2+ the intracellular Ca2+ stores were depleted by inhibition of the vesicular Ca2+ pump by thapsigargin (1 μM, Molecular Probes).
Experiments were performed with Ringer solution containing: 125 mM/l NaCl, 5 mM/l KCl, 1.2 mM/l MgSO4, 2 mM/l CaCl2, 2 mM/l Na2HPO4, 32 mM/l HEPES, and 5 mM/l glucose (pH 7.4). Nominally Ca2+-free solutions contained: 125 mM/l NaCl, 5 mM/l KCl, 1.2 mM/l MgSO4, 2 mM/l Na2HPO4, 32 mM/l HEPES, 0.5 mM/l EGTA, and 5 mM/l glucose (pH 7.4). For calibration purposes ionomycin (10 μM, Sigma-Aldrich) was applied at the end of each experiment.
Patch clamp experiments were performed at room temperature in voltage-clamp, fast-whole-cell mode according to Hamill et al. (18). The cells were continuously superfused through a flow system inserted into the dish. The bath was grounded via a bridge filled with NaCl Ringer solution. Borosilicate glass pipettes (1–3 MOhm tip resistance; GC 150 TF-10, Clark Medical Instruments) manufactured by a microprocessor-driven DMZ puller (Zeitz) were used in combination with a MS314 electrical micromanipulator (MW, Märzhäuser). The currents were recorded by an EPC-9 amplifier (Heka) using Pulse software (Heka) and an ITC-16 Interface (Instrutech). For ICRAC measurements whole-cell currents were elicited by 200 ms square wave voltage pulses from −100 to + 80 mV in 20 mV steps delivered from a holding potential of 0 mV. Alternatively, the currents were recorded with 200 ms voltage ramps from −120 to +100 mV. Kv whole-cell currents were elicited by 200 ms square wave voltage pulses from −90 to + 90 mV in 20 mV steps delivered at 20 ms intervals from a holding potential of −70 mV. The currents were recorded with an acquisition frequency of 10 and 3 kHz low-pass filtered.
For ICRAC measurements, cells were superfused with a bath solution containing: 140 mM/l NaCl, 5 mM/l KCl, 20 mM/l glucose, 10 mM/l HEPES/NaOH (pH 7.4), and the indicated concentration of divalent cations (0, 1, or 10 mM/l CaCl2 and 0, 1, or 10 mM/l MgCl2). A divalent-free bath solution contained 0.5 mM/l EGTA. In some experiments, extracellular Na+ was substituted by NMDG+ and the bath solution contained: 145 mM/l NMDG-Cl, 10 mM/l HEPES/NMDG, 20 mM/l glucose, 1 mM/l MgCl2, and 10 mM/l CaCl2. The patch clamp pipettes were filled with an internal solution (CsCl/NaCl pipette solution) containing: 120 mM/l CsCl, 35 mM/l NaCl, 1 mM/l MgATP, 10 mM/l EGTA, and 10 mM/l HEPES/CsOH (pH 7.4).
For Kv current measurements, the cells were superfused with a bath solution containing: 140 mM/l NaCl, 5 mM/l KCl, 1 mM/l MgCl2, 2 mM/l CaCl2, 20 mM/l glucose, and 10 mM/l HEPES/NaOH (pH 7.4). The patch clamp pipettes were filled with an internal solution (KCl/K-gluconate pipette solution) containing: 80 mM/l KCl, 60 mM/l K+-gluconate, 1 mM/l MgCl2, 1 mM/l Mg-ATP, 1 mM/l EGTA, 10 mM/l HEPES/KOH (pH 7.2).
Where indicated, SKF-96365 (10 μM, Sigma-Aldrich) or a combination of margatoxin (MgTx, 0.1 nM, Alomone Laboratories) and ICAGEN-4 (10 μM, Ref. 17) was added to the bath solution.
Total RNA was isolated from mouse DCs by using the Qiashredder and RNeasy Mini Kit from Qiagen. For cDNA first strand synthesis, 1 μg of total RNA in 12.5 μl of DEPC-H2O was mixed with 1 μl of oligo-dT primer (500 μg/ml, Invitrogen) and heated for 2 min at 70°C. A mix of 2 μl of 10× reaction buffer (Biolabs), 1 μl of dNTP mix (dATP, dCTP, dGTP, dTTP, 10 mM each, Promega), 0.5 μl of recombinant RNase inhibitor (Roche), 0.1 μl of M-MuLV reverse transcriptase (Biolabs), and 2.9 μl of DEPC-H2O was then added and the reaction mixture was incubated for 60 min at 42°C. The reaction was stopped by heating the mixture for 5 min at 94°C. The cDNA was stored at −80°C until PCR analysis. PCR analysis was then performed with 1 μl of the reverse transcription product in a total volume of 25 μl of a PCR mix containing 22 μl of sterile bi-distilled H2O, 1 μl of sense primer (100 pmol/μl), 1 μl of antisense primer (100 pmol/μl), and 1 puReTaq Ready-To-Go PCR bead (Amersham Biosciences) through 40 cycles (30 s at 95°C; 20 s at 58°C (CRACM1), at 56°C (CRACM2), or at 52°C (CRACM3); 45 s at 72°C). The following primers were used to amplify specific cDNA fragments from mouse DCs: Mouse CRACM1: sense primer: 5′-CATGGTAGCGATGGTGGAAGTC-3′; antisense primer: 5′-TGCTCATCGTCTTTAGTGCCT-3′. Mouse CRACM2: sense primer: 5′-ATGGTGGCCATGGTGGAGGT-3′; antisense primer: 5′-ATTGCCTTCAGCGCCTGCA-3′. Mouse CRACM3: sense primer: 5′-AAGCTCAAAGCCTCCAGCCGC-3′; antisense primer: 5′-GGTGGGTATTCATGATCGTTCT-3′. PCR products were analyzed by agarose gel electrophoresis.
TNF-α and IL-6 concentrations in DC culture supernatants were determined by using OptEIA ELISA kit (BD Pharmingen) according to the manufacturer’s protocol.
DC phagocytosis assay
DCs (106 cells/ml) were suspended in prewarmed serum-free RPMI 1640 medium, pulsed with FITC-conjugated dextran (Sigma-Aldrich) at a final concentration of 1 mg/ml, and incubated for 3 h at 37°C. Uptake of FITC-conjugated dextran was stopped by adding ice-cold PBS. The cells were then washed three times with ice cold PBS supplemented with 5% FCS and 0.01% sodium azide before FACS analysis.
DC migration assay
DCs were washed twice with PBS and resuspended in RPMI 1640 medium. Migration was assessed in triplicate in a multiwell chamber with 8-μm pore size filter (Calbiochem). The cell suspension (5 × 105 cells/ml) was placed in the upper chamber to migrate into the lower chamber in which either CCL21 (250 ng/ml, PeproTech) or medium alone as a control for spontaneous migration were included. The chamber was placed in a 5% CO2 37°C incubator for 4 h. The cells that migrated into the lower chamber were detached using Cell Detachment Buffer containing Calcein-AM fluorescent dye. The results were read using a standard fluorescence plate reader. The mean fluorescence of spontaneously migrated cells was subtracted from the total fluorescence of migrated cells.
Data are provided as means ± SEM, n represents the number of independent experiments. Differences were tested for significance using Student’s unpaired and paired two-tailed t test or ANOVA. p < 0.05 was considered statistically significant.
Stimulation of DCs with LPS (100 ng/ml) resulted in a rapid increase in [Ca2+]i (Fig. 1,A). This increase was due to Ca2+ release from intracellular stores and influx of extracellular Ca2+. Accordingly, removal of extracellular Ca2+ (Fig. 1,B) or the presence of SKF-96365, a known inhibitor of store-operated channels (10 μM, Fig. 1, A and C), significantly blunted but did not fully abrogate the increase of [Ca2+]i following LPS treatment. Store-operated Ca2+ entry in mouse DCs could be routinely measured upon store depletion by inhibition of the vesicular Ca2+-ATPase by thapsigargin (Fig. 1 D) and thus, we hypothesized that LPS treatment may lead to activation of CRAC channels.
To prove this hypothesis, whole-cell voltage clamp experiments were performed to study the entry of extracellular Ca2+ upon LPS stimulation of DCs. Within 1.5–3 min, LPS addition activated an inward current with properties similar to ICRAC (Fig. 2, A and B). With Ca2+ as charge carrier (10 mM Ca2+ in the bath) the LPS-stimulated current reversed at > +50 mV (Fig. 2,B), indicating high selectivity for Ca2+. In LPS-stimulated cells neither reversal potential of the current/voltage (I/V) relationship nor current amplitude were altered by replacing extracellular Na+ by NMDG+ (Fig. 2,A). The I/V curve of the LPS-stimulated current fraction revealed a prominent inward rectification at negative voltages. Reducing the concentration of external Ca2+ in the continued presence of external Na+ and Mg2+ reduced the inward current (Fig. 2, C, D, and F). One of the distinguishable features of CRAC channels is anomalous mole fraction, describing that when external Ca2+ concentration is very low (submicromolar range), large Na+ currents readily flow through the channels (19). In a Na+-containing but divalent cation-free external solution, the LPS-stimulated channel exhibited anomalous mole fraction becoming permeable for Na+ (Fig. 2, E and F). In the presence of extracellular Ca2+, the inward current of LPS-stimulated cells inactivated fast during hyperpolarizing voltages pulses (Fig. 2,F). This inactivation was absent when the cells were recorded in a divalent cation-free solution (Fig. 2,F). The current was inhibited by 10 μM SKF-96365, a blocker of store-operated channels (Fig. 2 G).
The membrane potential of DCs was measured using the current-clamp mode of the patch-clamp technique. As shown in Fig. 2 H, application of LPS led to membrane depolarization from −19.5 ± 2.5 mV (n = 8) to +1.8 ± 3.5 mV (n = 8).
Recently, the proteins involved in store-operated Ca2+ entry have been identified. These are STIM1, a Ca2+ sensor in the endoplasmic reticulum (20, 21, 22), and Orai 1 (or CRACM1), a pore subunit of the CRAC channel (23, 24, 25). Moreover, there are three mammalian homologous CRAC channel proteins, CRACM1, CRACM2, and CRACM3 (26). To test which CRAC channels are expressed in mouse DCs, DNA fragments specific for the cloned mouse CRACM1, CRACM2, and CRACM3 channels were amplified by RT-PCR. The RT-PCR data demonstrated endogenous expression of all three channels in mouse DCs (Fig. 2 I).
DCs are known to express voltage-gated K+ channels belonging to the Shaker (Kv1) family, presumably Kv1.3 and Kv1.5 (13, 14). Accordingly, margatoxin (MgTx, 0.1 nM) and ICAGEN-4 (10 μM, Ref. 17), blockers of Kv1.3 and Kv1.5, respectively, inhibited Kv-like currents in DCs (Fig. 3,A). The current fraction sensitive to MgTx (0.1 nM) alone was 54.1 ± 11.8% (n = 4, calculated for the current at + 60 mV), to ICAGEN-4 (10 μM) alone 65.7 ± 3.4% (n = 8), and when MgTx (0.1 nM) and ICAGEN-4 (10 μM) were applied together 84.2 ± 3.4% (n = 4) of the current was inhibited (the amplitude of the remaining current at +60 mV was significantly (p = 0.03) smaller than under control conditions). To test whether these Kv channels can modulate the Ca2+ entry through CRAC, the influence of MgTx and ICAGEN-4 on LPS-induced increase in [Ca2+]i has been determined using fura-2 Ca2+-imaging. As a result MgTx (1 nM) and ICAGEN-4 (10 μM) significantly reduced the LPS-induced rise in [Ca2+]i (Fig. 3, B and C).
TNF-α production in LPS-stimulated macrophages has previously been shown to depend on a transient increase in [Ca2+]i (10). Therefore, we examined the effect of channel blockers on TNF-α production. To this end, DCs were stimulated for 4 h with LPS (100 ng/ml) in the presence or absence of SKF-96365. As a result, treatment with SKF-96365 (10 μM, 30 min before addition of LPS and then during LPS stimulation) significantly blunted the TNF-α release from LPS stimulated DCs (Fig. 4,A). Similarly, MgTx (1 nM) and ICAGEN-4 (10 μM) significantly reduced the TNF-α release following LPS stimulation (Fig. 5,A). Moreover, production of IL-6 was as well significantly reduced by these channel blockers (Figs. 4,B and 5 B).
Next, we determined whether the maturation and activation of DCs incubated with LPS (100 ng/ml, 48 h) was different in the absence and in the presence of SKF-96365 (10 μM) or MgTx (1 nM) and ICAGEN-4 (10 μM). First, cells were collected and stained for MHC class II, CD86, and CD54. The CD11c+ gated population was analyzed for the expression of the mentioned markers. There was no significant difference in the expression of CD86 and CD54 before and after activation of LPS in the absence or in the presence of the channel blockers. However, the up-regulation of MHC class II by LPS was significantly reduced by SKF-96365 and MgTx plus ICAGEN-4 (Fig. 6 A), indicating that Ca2+ entry can be involved in DC maturation.
At the same time, the phagocytic capacity of DCs assessed as FITC-dextran uptake, was dramatically increased by SKF-96365 or MgTx and ICAGEN-4 (Fig. 6 B), suggesting that inhibition of CRAC and Kv channels leads to a less mature DC phenotype with higher phagocytic activity.
Ability of DCs to migrate to lymphoid tissues is fundamental for the launching and the coordination of immune responses (27). Therefore, we determined whether DC migration in response to a CCR7 ligand CCL21 is influenced by CRAC and Kv channel blockers. Treatment with SKF-96365 or with MgTx and ICAGEN-4 led to a significant impairment of the ability of LPS-stimulated DCs to migrate in response to CCL21 (Fig. 7).
In the present study, we demonstrated that LPS causes a rapid transient increase of cytosolic Ca2+ activity ([Ca2+]i) in mouse bone marrow-derived DCs. Removal of extracellular Ca2+ or application of SKF-96365, a blocker of store-operated calcium channels (ICRAC) (19), blunted, but did not completely abolish peak and slope of [Ca2+]i increase. Thus, LPS-induced increase in [Ca2+]i was partially due to Ca2+ influx through Ca2+ channels in the plasma membrane and partially due to Ca2+ release from intracellular stores. ICRAC has been demonstrated to be the principal Ca2+ entry pathway in DCs (12). ICRAC could be activated by store depletion induced by dialyzing the cytosol with inositol 1,4,5-trisphosphate and with high concentration of the Ca2+ chelator BAPTA, which chelates Ca2+ that leaks from the stores and hence prevents store refilling (12). Our experiments reveal that LPS strongly activates ICRAC and thus disclose an important mechanism of channel activation.
We further show that the LPS induced [Ca2+]i increase in DCs is important for subsequent TNF-α and IL-6 production. Accordingly, inhibition of ICRAC impairs TNF-α and IL-6 secretion. Moreover, LPS-induced up-regulation of MHC class II expression is impaired and phagocytic capacity enhanced in DCs matured in the presence of SKF-96365. In addition, DC migration is dependent on ICRAC activity. Our study thus demonstrates a pivotal role of ICRAC for DC maturation and activity.
ICRAC has been suggested to be important for DC maturation, as activation of ICRAC with thapsigargin induced marked maturation of mouse myeloid DCs (12), human peripheral blood monocytes, and HL-60 cells (3). Also the addition of calcium ionophore to human monocytes or immature DCs resulted in the acquisition of many properties characteristic of activated myeloid DCs (4). However, physiological Ca2+ signaling occurs through Ca2+ oscillations and not through a long-lasting increase or decrease in [Ca2+]i and the activity of signal terminators, such as ER- and plasma membrane-localized Ca2+ pumps (SERCA and PMCA, respectively), plasma membrane exchangers (Na+-Ca2+ exchanger), mitochondrial and cytosolic buffer proteins is extremely important for determination of duration, amplitude, and intracellular location of a particular Ca2+ signal (28).
It is important to note that SKF-96365 which is known to inhibit store-operated Ca2+ channels, appears to be equally potent in blocking voltage-dependent Ca2+ and TRP channels (19) and, thus, cannot be considered as a specific CRAC channel blocker. Recently, the molecules involved in store operated Ca2+ entry have been identified. These are STIM1, a Ca2+ sensor in the endoplasmic reticulum (20, 21, 22), and Orai 1–3 (or CRACM1–3), pore-forming subunits of CRAC channel (23, 24, 25). The identification of STIM1 and Orai should allow development of potent and specific inhibitors of CRAC channel in near future.
Entry of positively charged Ca2+ through ICRAC is expected to be a function of cell membrane potential and thus K+ channel activity. Thus, we explored the influence of Kv channels on Ca2+ entry. As a result, we indeed observed that Kv channels in DCs sustain the increase of [Ca2+]i upon LPS stimulation. They are most probably effective by maintaining the negative membrane potential (29) and providing the necessary electrical driving force for Ca2+ influx through ICRAC. Recently, Kv1.3 and Kv1.5 channels were discovered in human blood-derived DCs (13) and bone marrow derived DCs from mice (14). The channels seem to play a role in the maturation process of DCs, because costimulatory molecule expression and cytokine production is decreased in DCs matured in the presence of Kv channel blockers (13, 14). The present study shows that, similar to CRAC blocker SKF-96365, Kv channel blockers led to impaired TNF-α and IL-6 production, decreased LPS-induced up-regulation of MHC II expression, enhanced phagocytic capacity, and reduced migration in mouse DCs and thus provides evidence for novel Kv channel-dependent DC functions. Kv channels are similarly involved in the activation and proliferation of leukocytes (15). Kv1.3 constitutes the dominant K+ conductance of resting T lymphocytes (30). Inhibition of Kv1.3 channels induces membrane depolarization and prevents the activation response of human T cells (15). Moreover, Kv channels are regulated during proliferation and activation of macrophages (31).
In summary, this study shows that DCs respond to LPS stimulation with a fast increase of [Ca2+]i, which is accomplished by both, Ca2+ release from intracellular stores and Ca2+ influx through ICRAC. The Ca2+ influx through ICRAC depends on the activity of Kv channels. Inhibition of either ICRAC or Kv channels leads to profound changes in DC functions, including changes in maturation, phagocytosis, migration, and cytokine production.
We gratefully acknowledge the expert technical assistance by Andrea Janessa and the meticulous preparation of the manuscript by Lejla Subasic. We thank Dr. S. Huber and Dr. O. Ureche for helpful discussion.
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.
This work was supported by by the Deutsche Forschungsgemeinschaft DFG (SFB 766) and The International Graduate School (GRK 1302/1) “The PI3K Pathway in Tumor Growth and Diabetes”.
Abbreviations used in this paper: DC, dendritic cell; CamK, Calmodulin kinase; CRAC, Ca2+ release-activated Ca2+ channel; Kv, voltage-gated K+; MgTx, margatoxin.