Blunted IgE-Mediated Activation of Mast Cells in Mice Lacking the Ca²⁺-Activated K⁺ Channel K_Ca3.1¹ Free

Allergic reactions of the immediate type (1, 2), including allergic rhinitis (3), asthma (4), and anaphylaxis, are mediated by the cross-linking of IgE receptors (FcεRI) on mast cells. In addition, mast cells contribute to the development of T cell-mediated delayed-type hypersensitivity reactions such as contact hypersensitivity or experimental allergic encephalitis, a murine model of multiple sclerosis (5, 6, 7). Upon activation, mast cells release a variety of factors such as cytokines and proteases and thus regulate the function of inflammatory cells such as neutrophils and T cells (5, 8, 9, 10, 11, 12, 13, 14, 15, 16). Moreover, it is well-understood that mast cells, similar to dendritic cells, can serve as important mediators between innate and adaptive immunity (17).

Mast cells can secrete histamine and immunoreactive leukotriene C₄ in response to nonimmunological stimuli, such as endothelin-1, a potent vasoconstrictor (18, 19). Mast cell activation, upon both FcεRI aggregation and endothelin-1 stimulation, involves an increase in intracellular Ca²⁺ concentration, which is partially accomplished by Ca²⁺ release from intracellular stores (20, 21, 22, 23, 24, 25). The increase of cytosolic Ca²⁺ also results from activation of store-operated Ca²⁺ channels (SOCs)⁵ in the plasma membrane, which are important for the release of inflammatory mediators (26, 27, 28, 29, 30, 31, 32, 33).

The electrical driving force for Ca²⁺ entry is provided by the cell membrane potential, which is maintained by the activity of K⁺ channels. Activation of K⁺ channels, with subsequent hyperpolarization of the cell membrane, enhances mast cell degranulation, because cell membrane Ca²⁺ influx is greater at negative membrane potentials (34). Conversely, depolarization of the cell membrane by activation of the cation channel TRPM4 decreases Ca²⁺ influx and subsequent degranulation (35). Furthermore, decreasing the chemical driving force for K⁺ exit by increasing extracellular K⁺ concentration (36) or inhibiting K⁺ channels with Ba²⁺ and quinidine (37) inhibits mast cell degranulation.

K⁺ channels expressed in mast cells include Ca²⁺-activated K⁺ channels, which have been implicated in the regulation of mast cell exocytosis (30). However, the nonselective K_Ca3.1 blocker charybdotoxin only modestly inhibited histamine release from human lung mast cells (38). In contrast, the K_Ca3.1 opener 1-EBIO enhanced IgE-dependent Ca²⁺ influx and degranulation in response to a submaximal stimulus (38). K_Ca3.1 has not been demonstrated in rodent mast cells by electrophysiological means. However, in the RBL-2H3 rat mast cell line charybdotoxin and the nonselective blocker cetiedil did attenuate degranulation and activation-dependent hyperpolarization of the plasma membrane (38, 39). However, another K_Ca3.1 blocker clotrimazole had no effect in these cells at pharmacologically relevant concentrations (39, 40). The precise role that K_Ca3.1 plays in mast cell mediator release is therefore unclear, although its presence does appear to be important for human lung mast cell migration (41). Moreover, the pharmacological tools are not necessarily specific for K_Ca3.1, because clotrimazole and TRAM-34 (1-[(2-chlorophenyl)diphenylmethyl]-1H-pyrazole) were reported to also inhibit nonselective cation channels (42, 43).

The present study has therefore been performed to define the contribution of the Ca²⁺-activated K⁺ channel K_Ca3.1 to mast cell function, both in vitro and in vivo. To this end, we isolated mast cells from gene-targeted mice lacking K_Ca3.1 (K_Ca3.1^−/−) (44) and their wild-type littermates (K_Ca3.1^+/+). Our findings show that loss of K_Ca3.1 channel expression significantly impairs mast cell volume regulation, IgE- and endothelin-1-induced mast cell Ca²⁺ influx and degranulation, as well as the anaphylactic response. K_Ca3.1 channels are, therefore, a critical regulator of mast cell function.

All animal experiments were conducted according to the guidelines of the American Physiological Society, as well as the German law for the welfare of animals, and were approved by local authorities. K_Ca3.1-deficient mice (K_Ca3.1^−/−) were generated as described (44) and maintained at the animal facility of the Pharmaceutical Institute (Department of Pharmacology and Toxicology, University of Tübingen, Tübingen, Germany). Briefly, targeted deletion of K_Ca3.1 in mice was performed using the Cre/loxP system yielding L1/+ clones. Correctly targeted L1/+ clones were injected into C57BL/6 blastocysts and the resulting chimeras were mated with 129Sv mice to obtain germline transmission. Heterozygous offspring were intercrossed with C57BL/6 mice to produce wild-type (K_Ca3.1^+/+) and K_Ca3.1^−/− mice on a hybrid 129Sv/C57BL/6 background (always F₂). Genotyping was performed by PCR using three primers amplifying either the wild-type (264 bp) or the knockout (507 bp) allele (for details, see Ref. 44). Homozygous K_Ca3.1^−/− mice are viable, fertile, and do not show reduced life expectancy. Either litter- or age-matched wild-type (K_Ca3.1^+/+) and K_Ca3.1^−/− mice of either sex (with hybrid 129Sv/C57BL/6 background, always F₂ generation) were randomly assigned to the experimental procedures with respect to German legislation on animal protection.

Mast cells were isolated from femoral bone marrow of 6- to 8-wk-old naive K_Ca3.1^+/+ and K_Ca3.1^−/− mice and cultured for 4 wk in RPMI 1640 (Invitrogen Life Technologies) containing 10% FCS, 1% penicillin/streptomycin, 20 ng/ml IL-3 (R&D Systems), and 100 ng/ml of the c-kit ligand stem cell factor (PeproTech). Mast cell numbers were determined by a trypan blue dye exclusion assay in 0.1% trypan blue. Viable (trypan blue-negative) cells were counted every 3–5 days and growth curves were drawn using the cultured time as abscissa and number of living cells as ordinate. BMMC maturation was confirmed by flow cytometry (FACSCalibur; BD Biosciences) using the following specific fluorescent-labeled Abs: PE-labeled anti-FcεRI (eBioscience), allophycocyanin-labeled anti-CD117 (BD Pharmingen), and FITC-labeled anti-CD34 (BD Pharmingen). Cells were kept in culture 4–6 wk before the experiments. For experiments, BMMCs were sensitized for 1 h with monoclonal mouse anti-DNP mouse IgE (anti-DNP IgE, 5–10 μg/ml per 1 × 10⁶ cells, clone SPE-7; Sigma-Aldrich) in culture medium and challenged with DNP-human serum albumin (DNP-HSA; 5–100 ng/ml; Sigma-Aldrich). Alternatively, BMMCs were stimulated by endothelin-1 (100 nM; PeptaNova).

Patch-clamp experiments have been performed at room temperature in voltage-clamp, fast whole-cell mode (45). BMMCs were continuously superfused by a flow system inserted into the dish. The bath was grounded via a bridge filled with NaCl Ringer solution, containing: 145 mM NaCl, 5 mM KCl, 2 mM MgCl₂, 1 mM CaCl₂, 10 mM glucose, 10 mM HEPES/NaOH (pH 7.4). Borosilicate glass pipettes (2- to 4-megaohm (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). Whole-cell currents were determined as 10 successive 200-ms square pulses from a −35 mV holding potential to potentials between −115 mV and +65 mV. The currents were recorded with an acquisition frequency of 10 and 3 kHz low-pass filtered.

The pipette solution contained: 140 mM K-gluconate, 5 mM KCl, 1.2 mM MgCl₂, 2 mM EGTA, 1.26 mM CaCl₂ (pCa 7), 2 mM Na₂ATP, and 10 mM HEPES/KOH (pH 7.2), and was used in combination with NaCl Ringer bath solution. Where indicated, the Ag DNP-HSA (50 ng/ml), endothelin-1 (100 nM), the channel blocker TRAM-34 (300 nM; Sigma-Aldrich), and/or the Ca²⁺ ionophore ionomycin (1 μM; Sigma-Aldrich) were added to the bath solution.

The offset potentials between both electrodes were zeroed before sealing. The potentials were corrected for liquid junction potentials as estimated according to Barry and Lynch (46). The original whole-cell current traces are depicted without further filtering and currents of the individual voltage square pulses are superimposed. The applied voltages refer to the cytoplasmic face of the membrane with respect to the extracellular space. The inward currents, defined as flow of positive charge from the extracellular to the cytoplasmic membrane face, are negative currents and are depicted as downward deflections of the original current traces.

Intracellular Ca²⁺ measurements were performed as described (47) using fura-2AM. Briefly, BMMCs were loaded with fura-2AM (2 μM; Molecular Probes) for 20 min at 37°C. The acetomethyl ester of fura-2 is an uncharged nonfluorescent molecule easily permeating the cell membrane. Once the dye is inside the cell, the lipophilic groups are cleaved by esterases. This results in the charged fluorescent form of the dye that is trapped within the cell. Additionally, the hydrolysis of the esterified groups is essential for binding of the target ion Ca²⁺. The cells were continuously superfused to remove leaked-out extracellular fura-2.

Fluorescence measurements were conducted with an inverted phase-contrast microscope (Axiovert 100; Zeiss). Cells were excited alternatively at 340 and 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) mounted on the microscope. Emitted fluorescence intensity was recorded at 510 nm and data acquisition was performed by using Metafluor computer software (Universal Imaging). To determine the fluorescence intensity in single cells, the regions of interest were determined as circuits closely surrounding individual cells.

After a control period to wash the cells and remove any nongathered dye, intracellular Ca²⁺ was measured before and following addition of DNP-HSA (50 ng/ml) to IgE-sensitized BMMCs, in the absence or presence of extracellular Ca²⁺. Alternatively, Ca²⁺ was measured upon stimulation of BMMCs with endothelin-1 (100 nM). As a measure of the increase of cytosolic Ca²⁺, the slope and peak of the changes in the 340/380-nm ratio were calculated for each experiment.

Experiments were performed with Ringer solution containing: 125 mM NaCl, 5 mM KCl, 1.2 mM MgSO₄, 2 mM CaCl₂, 2 mM Na₂HPO₄, 32 mM HEPES, 5 mM glucose (pH 7.4). To reach nominally Ca²⁺-free conditions, experiments were performed using Ca²⁺-free Ringer solution containing: 125 mM NaCl, 5 mM KCl, 1.2 mM MgSO₄, 2 mM Na₂HPO₄, 32 mM HEPES, 0.5 mM EGTA, 5 mM glucose (pH 7.4). For intracellular calibration purposes, ionomycin (10 μM) was applied at the end of each experiment.

Regulatory cell volume decrease was estimated from fura-2AM fluorescence at the intracellular Ca²⁺ concentration-insensitive isosbestic point (48). Briefly, the BMMCs were loaded with 2 μM fura-2AM for 20 min at 37°C and then excited at 357 nm. Fluorescence emission was recorded at 510 nm. To measure cell volume-regulatory behavior, the initial average fluorescence of the fura-2AM-loaded cells was obtained in standard Ringer (containing: 60 mM NaCl, 5 mM KCl, 1 mM CaCl₂, 1.2 mM MgSO₄, 2 mM Na₂HPO₄, 32 mM HEPES, 5 mM glucose (pH 7.4), 290 milliosmole (mosm)/L) for 2–3 min and was standardized to 1.00 (f₀). Subsequently, the osmolarity was reduced to 160 mosm/L and changes in the fluorescence intensity (f) according to changes in cell volume were recorded. For data interpretation, the measured fluorescence was compared with the initial fluorescence intensity (f/f₀).

Mature bone marrow mast cells were seeded on 96-well plates in fresh medium with anti-DNP IgE for 1 h. Afterward, cells were washed in Tyrode’s salt solution (Sigma-Aldrich) and challenged with DNP-HSA (50 ng/ml), or for positive control with 100 ng/ml PMA ester (Sigma-Aldrich) and 1 μM ionomycin (Sigma-Aldrich), for 15 min. A total of 20 μl of supernatant and 20 μl of 2 mM 4-nitrophenyl N-acetyl-β-d-glucosaminide (Sigma-Aldrich) diluted in 0.2 M citrate buffer (pH 4.5) was added to each well of the 96-well plate and color was developed for 2 h at 37°C. The reaction was terminated with 1 M Tris buffer (pH 9.0) and the absorbance was measured at 405 nm in an ELISA microplate reader (Sunrise remote; TECAN). The data were corrected for spontaneous release and are expressed as the percent of total release (0.1% Triton X-100). Histamine and IL-6 release from mast cells was measured by ELISA according to the instructions of the manufacturer (IBL and BD Biosciences/BD Pharmingen, respectively).

Anesthetized mice were euthanized by cervical dislocation and the abdominal skin was cleansed with 70% ethanol. A total of 4 ml of sterile 0.9% NaCl was then instilled into the peritoneum. The abdomen was massaged gently for 1 min and then opened with sterile scissors. Recovered peritoneal lavage fluid was centrifuged at 200 × g for 5 min and the cell pellets were resuspended in RBC lysis buffer for 1 min, recentrifuged, and then the cell pellet was resuspended in PBS. Slides were prepared as “thick blood films” and fixed with 4% paraformaldehyde. After staining with toluidine blue, at least 600 cells/slide were counted. Mast cell numbers are expressed as normalized to the corresponding leukocyte counts.

Anesthetized mice were euthanized by cervical dislocation and the skin was cleansed with 70% ethanol. Ears were cut off at the base, fixed in 4% paraformaldehyde overnight and finally embedded in paraffin. Three- to 4-μm-thick tissue sections of the midsections of the ears were prepared, deparaffinized, and stained with toluidine blue. Mast cell numbers of at least three slices per ear were determined using a Zeiss Axiovert 200 microscope with a LD Achroplan ×40 lens in bright field mode.

Mice were sensitized with 50 μg/250 μl of anti-DNP IgE by i.p. application. The next day, mice were challenged with either DNP-HSA (120 μg/200 μl) or PBS. Body temperature was monitored before and each 7 min after Ag challenge with a BAT-10 type T thermocouple thermometer and a RET-3 rectal probe for mice (Physitemp Instruments; distributed by World Precision Instruments) using a Duo18 data recording system (World Precision Instruments) during the midportion of the light phase of the light cycle. Mice were placed with the tail raised and the Vaseline-covered probe was inserted a standardized distance of 2 cm until a stable temperature reading was obtained. Baseline temperature was measured after habituating mice to rectal probe insertion. Ambient room temperature was 23°C and the animals were exposed to a 12-h light and 12-h dark cycle (7 a.m. to 7 p.m.). Data are expressed as a change in body temperature following treatment (Δ°C).

Data are provided as means ± SEM; n represents the number of animals/independent experiments. All data were tested for significance using the Student unpaired two-tailed t test, one sample t test, or ANOVA (Dunnett’s test), where applicable. A value of p < 0.05 was considered to indicate statistical significance.

To investigate the functional role of K_Ca3.1 in mast cells, cells were derived from the bone marrow (BMMCs) of K_Ca3.1^+/+ and K_Ca3.1^−/− mice; growth (Fig. 1,A) and expression of the mast cell surface markers CD117, CD34, and FcεRI (Fig. 1, B and C) were determined. No significant difference in the cell growth or the abundance of any of the three markers for mast cell maturation was observed between BMMCs of the two genotypes (Fig. 1, A and C). Thus, targeted deletion of K_Ca3.1 in mice did not influence the growth and maturation of BMMCs.

The density of mast cells in skin, analyzed by staining of ear sections with toluidine blue, was comparable in K_Ca3.1^+/+ and K_Ca3.1^−/− mice (Fig. 1,D). Similarly, the number of mast cells in the peritoneum was not different between genotypes (Fig. 1 E).

To analyze the currents upon stimulation of mast cells, BMMCs were incubated with IgE (5 μg/ml) directed against the hapten DNP for 1 h, washed and then stimulated with Ag, DNP coupled to the carrier protein HSA (DNP-HSA, 50 ng/ml). According to patch-clamp experiments, the exposure of BMMCs to Ag was followed by a sharp increase of K⁺-selective conductance in K_Ca3.1^+/+ but not in K_Ca3.1^−/− BMMCs (Fig. 2, A–D). The K⁺ current was inhibited by TRAM-34, but not by apamin or iberiotoxin (Fig. 2,B and data not shown). Similarly, treatment of the cells with ionomycin led to activation of the K⁺ channels in K_Ca3.1^+/+ BMMCs but not in K_Ca3.1^−/− BMMCs (Fig. 2 D). Thus, K_Ca3.1 accounts for Ca²⁺-activated K⁺ channel activity in BMMCs.

The membrane potential of BMMCs was measured using the current-clamp mode of the patch-clamp technique. Before application of DNP-HSA to sensitized BMMCs, the membrane potential was more depolarized (−17.0 ± 0.6 mV, n = 7) in K_Ca3.1^−/− than in K_Ca3.1^+/+ (−35.6 ± 5.3 mV, n = 10) BMMCs (Fig. 2 E). Applying TRAM-34 (300 nM) under resting conditions led to a pronounced depolarization (−15.4 ± 1.9 mV, n = 5) in K_Ca3.1^+/+ cells, demonstrating dependence of the resting cell membrane potential on K_Ca3.1. Stimulation with 50 ng/ml DNP-HSA, was followed by a hyperpolarization to −56.9 ± 4.6 mV (n = 10) in K_Ca3.1^+/+ cells, but did not significantly alter the membrane potential of K_Ca3.1^−/− BMMCs (−17.4 ± 1.1 mV, n = 7). The Ca²⁺ ionophore ionomycin (1 μM) similarly hyperpolarized K_Ca3.1^+/+ cells to −59.0 ± 5.9 mV (n = 5), but had no significant effect on the membrane potential of K_Ca3.1^−/− BMMCs (−18.8 ± 1.8 mV, n = 5). In K_Ca3.1^+/+ cells, inhibition of DNP-HSA-induced current with TRAM-34 reversed the hyper polarization (−24.4 ± 2.6 mV, n = 12). Thus, K_Ca3.1 plays an important role in maintaining the resting membrane potential and in hyperpolarization upon FcεRI stimulation.

Osmotic swelling of most cells is followed by regulatory cell volume decrease (RVD), which depends on K⁺ channel activity in a wide variety of cells (49). RVD was indeed impaired in K_Ca3.1^−/− BMMCs (Fig. 3,A). The slope and Δ of the recovery was decreased in K_Ca3.1^−/− as compared with K_Ca3.1^+/+ BMMCs (Fig. 3 B).

According to fura-2 fluorescence experiments, exposure of IgE-sensitized mast cells to Ag led to a sharp increase in cytosolic Ca²⁺ concentration in K_Ca3.1^+/+ BMMCs. The Ca²⁺ increase following Ag stimulation was markedly reduced in K_Ca3.1^−/− BMMCs (Fig. 4,A). Correspondingly, the maximal fluorescence Δratio (peak) and the slope of the ratio (Δratio/time) upon stimulation with Ag were significantly smaller in K_Ca3.1^−/− than in K_Ca3.1^+/+ BMMCs (Fig. 4,B). Exposure of BMMCs to a Ca²⁺-free solution was followed by a decline of cytosolic Ca²⁺ concentration reflecting extrusion of Ca²⁺ by transport systems such as Na⁺/Ca²⁺ exchanger (Fig. 4,C). Stimulation of the BMMCs with Ag in Ca²⁺-free solution allowed an estimate of Ca²⁺ release from intracellular stores (Fig. 4,C). This release was not significantly different between the genotypes (Fig. 4 D).

The critical role of K_Ca3.1 in SOC-mediated Ca²⁺ influx into BMMCs suggests an important contribution of this K⁺ channel type for mast cell function, especially for Ca²⁺-dependent processes, such as degranulation. To test this hypothesis, we analyzed the ability of K_Ca3.1^+/+ and K_Ca3.1^−/− BMMCs to undergo degranulation in response to IgE-induced stimulation. Upon stimulation, BMMCs release β-hexosaminidase, an enzyme stored in mast cell granules (50). The granule content determined by total β-hexosaminidase levels after cell lysis with Triton X-100 was indistinguishable between the two genotypes (p = 0.80, t test). However, IgE-dependent release of β-hexosaminidase was significantly smaller in K_Ca3.1^−/− as compared with K_Ca3.1^+/+ BMMCs, upon stimulation with Ag in a wide range of concentrations (5–100 ng/ml; Fig. 5,A). Similarly, K_Ca3.1^−/− BMMCs released less β-hexosaminidase when stimulated with 1 μM ionomycin and 100 ng/ml PMA (Fig. 5 B). The production of IL-6 was not significantly blunted in K_Ca3.1^−/− BMMCs, though there was a downward trend (2172.34 ± 138.36 pg/ml, n = 3 in K_Ca3.1^+/+ and 1655.98 ± 24.08 pg/ml, n = 3 in K_Ca3.1^−/− BMMCs after 24 h following FcεRI stimulation). Thus, K_Ca3.1 is indeed a decisive regulator of mast cell degranulation.

To evaluate whether the observed alterations in K_Ca3.1^−/− BMMCs were also relevant for mast cell function in vivo, we triggered passive systemic anaphylaxis in mice. K_Ca3.1^+/+ and K_Ca3.1^−/− mice were sensitized with anti-DNP IgE i.p. After overnight rest, mice received DNP-HSA Ag or saline as a control by i.p. injection and body temperature was monitored over time. As shown in Fig. 6, the decrease in body temperature following Ag treatment was markedly attenuated in K_Ca3.1^−/− mice, an observation pointing to an impaired in vivo function of K_Ca3.1-deficient mast cells.

BMMCs express endothelin A receptors, which are associated with Ca²⁺-dependent release of histamine and leukotriene C₄ (18, 19). Additional experiments were thus performed to test whether K_Ca3.1 is similarly involved in the response to endothelin-1. Exposure of BMMCs to endothelin-1 (100 nM) was indeed followed by an increase of K⁺-selective conductance and hyperpolarization in K_Ca3.1^+/+, but not in K_Ca3.1^−/−, BMMCs (Fig. 7, A–C). The Ca²⁺ entry upon endothelin-1 stimulation was significantly reduced in K_Ca3.1^−/− BMMCs (Fig. 7, D and E). Moreover, K_Ca3.1^−/− cells secreted significantly less histamine as compared with K_Ca3.1^+/+ BMMCs upon endothelin-1 stimulation (Fig. 7 F). Thus, K_Ca3.1 is similarly involved in the effects of endothelin-1 on Ca²⁺-dependent mast cell responses.

The present study discloses a key role for K_Ca3.1 in IgE-dependent mast cell degranulation. Targeted deletion of K_Ca3.1 in mice resulted in mast cell hyporesponsiveness and, as a consequence, in marked attenuation of the IgE-dependent systemic anaphylactic response. The presence of K_Ca3.1 in mast cells has been demonstrated previously in human lung mast cells by patch-clamp electrophysiology and by the presence of K_Ca3.1 mRNA in these cells (29, 30, 38). K_Ca3.1 opening is not required for, but potentiates, mast cell secretion (29, 30, 38) and plays a vital role in cell migration (41). The effect of drugs able to block K_Ca3.1 also suggested that the channel was present in the RBL-2H3 mast cell line (39, 40). The present study demonstrates for the first time that mouse BMMCs indeed express K_Ca3.1, and that BMMCs derived from K_Ca3.1-deficient mice completely lack Ca²⁺-activated K⁺ channel activity. We show that loss of K_Ca3.1 in mouse BMMCs markedly attenuates their IgE-dependent degranulation both in vitro and in vivo.

Our findings confirm previous work that K_Ca3.1 plays a critical role in the regulation of mast cell membrane potential, and thus influences the influx of extracellular Ca²⁺. The initial increase in cytoplasmic [Ca²⁺] following FcεRI stimulation results in the activation of K_Ca3.1 channels, which hyperpolarize the cell membrane and thus provide the electrical driving force for Ca²⁺ influx through store-operated Ca²⁺ channels. Moreover, hyperpolarization increases the open probability of the Ca²⁺ channels, which are inwardly rectifying (34). Recently, it was shown that the Ca²⁺-activated nonselective cation channel TRPM4 in mast cells acts as a negative regulator of Ca²⁺ influx following FcεRI stimulation, because the entry of cations depolarizes the membrane and limits the electrical driving force for Ca²⁺ entry through SOCs (35) and/or nonspecific cation channels permeable to divalent cations (51, 52, 53). Thus, the Ca²⁺-activated K_Ca3.1 channel provides positive feedback for the Ca²⁺ signal, whereas the Ca²⁺-activated TRPM4 acts as a molecular “brake” on Ca²⁺ influx.

However, the relevance of K_Ca3.1 for mast cell secretion was not entirely clear from previous studies. This was likely because the use of pharmacological inhibitors is fraught with difficulties due to nonspecific effects. For example, clotrimazole and TRAM-34 are reported to inhibit nonselective cation channels (42, 43). Our study unequivocally shows that the regulation of membrane potential by K_Ca3.1 plays a critical role in regulating the magnitude of Ca²⁺ influx and in turn degranulation.

IgE-dependent degranulation is not the only K_Ca3.1-dependent process in mast cells. Stimulation of mast cells via endothelin receptors, which similarly requires Ca²⁺ entry (18, 19) was also impaired in K_Ca3.1^−/− BMMCs. Thus, the channel is important in diverse Ca²⁺-dependent processes of mast cells.

Targeted deletion of K_Ca3.1 in mice had no effect on mast cell growth and development. This is in good agreement with the data showing that blocking K_Ca3.1 was unable to inhibit human lung mast cell proliferation (41). We have demonstrated that the number of mast cells is normal in K_Ca3.1^−/− mice. This observation is also important because it demonstrates that targeting K_Ca3.1 may not impair beneficial mast cell functions such as host defense, which makes this channel an even more attractive pharmacological target.

Beyond their role in the regulation of mast cell function, Ca²⁺-activated K⁺ channels serve a multitude of further functions in other cells. Accordingly, several defective functions have previously been described in K_Ca3.1 knockout mice, including elevated blood pressure (54) and impaired T lymphocyte and erythrocyte volume regulation (55). We show here that RVD of BMMCs was also dependent on K_Ca3.1. The present novel observation of deranged mast cell function adds to the hitherto known defects of K_Ca3.1-deficient mice.

In conclusion, this is the first study to show that K_Ca3.1 is a critical regulator of mast cell degranulation, which plays an important role in allergic responses and anaphylaxis. Thus, K_Ca3.1 may represent a novel target for antiallergic therapy.

We thank Andrea Janessa and Clement Kabagema for excellent technical assistance. We gratefully acknowledge the meticulous preparation of the manuscript by Lejla Subasic and Tanja Loch. We thank Dr. S. Huber for helpful discussion and V. Shumilin for help during data analysis.

The authors have no financial conflict of interest.