Chloride ion efflux is an early event occurring after exposure of human neutrophils to several soluble agonists. Under these circumstances, a rapid and reversible fall in the high basal intracellular chloride (Cli) levels is observed. This event is thought to play a crucial role in the modulation of several critical neutrophil responses including activation and up-regulation of adhesion molecules, cell attachment and spreading, cytoplasmic alkalinization, and activation of the respiratory burst. At present, however, no data are available on chloride ion movements during neutrophil phagocytosis. In this study, we provide evidence that phagocytosis of Candida albicans opsonized with either whole serum, complement-derived opsonins, or purified human IgG elicits an early and long-lasting Cl efflux accompanied by a marked, irreversible loss of Cli. Simultaneous assessment of Cl efflux and phagocytosis in cytochalasin D-treated neutrophils indicated that Cl efflux occurs without particle ingestion. These results suggest that engagement of immune receptors is sufficient to promote chloride ion movements. Several structurally unrelated chloride channel blockers inhibited phagocytosis-induced Cl efflux as well as the release of azurophilic—but not specific—granules. It implicates that different neutrophil secretory compartments display distinct sensitivity to Cli modifications. Intriguingly, inhibitors of Cl exchange inhibited cytosolic Ca2+ elevation, whereas Cl efflux was not impaired in Ca2+-depleted neutrophils. We also show that FcγR(s)- and CR3/CR1-mediated Cl efflux appears to be dependent on protein tyrosine phosphorylation but independent of PI3K and phospholipase C activation.

Neutrophils use part of their energy to pump chloride ions from the extracellular environment into the intracellular compartment, thereby accumulating a considerable amount of this anion (1). Accordingly, a distinct feature of resting neutrophils is an unusually high (80–100 mM) intracellular Cl concentration that is 4- to 5-fold higher than predicted on the basis of the Nernst equation (1, 2). The discovery of high basal chloride ions content (Cli)2 (2) in human neutrophils dates back approximately 40 years ago (3), but information on the role of chloride in the neutrophil pathophysiology has only recently started to emerge. In this regard, a major breakthrough was made when it was shown that a dramatic loss of Cli occurs when neutrophils are exposed to various physiological and nonphysiological soluble agonists (4). Many groups, including our own, have contributed to identify causal relationships between changes in Cli and activation of several neutrophil responses. Accordingly, direct measurements of Cl movements in stimulated neutrophils have provided evidence that Cl fluxes are involved in the modulation of several critical neutrophil responses including activation and up-regulation of adhesion molecules, cell attachment and spreading, cytoplasmic alkalinization, and activation of the respiratory burst (5, 6, 7, 8, 9). These findings have led to the hypothesis that Cl may act as a second messenger in human neutrophils. At present, however, the mechanisms whereby Cli fluctuations regulate the complex events leading to neutrophil activation have not been elucidated.

Compared with other inflammatory cells, neutrophils have the largest repertoire of secretory compartments containing a number of proteins involved in cell adhesion, recognition of opsonized particles, generation of reactive oxygen species, and killing and degradation of microorganisms. In this regard, at least four distinct types of granules have been described within the cytoplasm, namely azurophilic or primary granules, specific or secondary granules, gelatinase-rich tertiary granules, and secretory vesicles (10, 11). These compartments are dispersed throughout the cytoplasm in unstimulated neutrophils. However, upon activation with phagocytosable or soluble agonists, granules fuse with the plasma membrane, thereby discharging their content from the cell (12, 13, 14, 15, 16, 17, 18, 19, 20). It has been previously demonstrated that secretory compartments are released following a precise hierarchy that largely depends on their distinct sensitivity to cytosolic Ca2+ concentrations. Specifically, it has been shown that secretory vesicles display the lowest Ca2+ threshold followed—in increasing order—by tertiary, specific, and azurophilic granules (21, 22).

Weissmann and coworkers first reported that anion channel blockers have the capacity to inhibit lysosomal enzyme secretion in human neutrophils stimulated by either immune complexes or the calcium ionophore A23187 (23, 24). Although these results pointed to a role of permeant anions in neutrophil degranulation, both the identity of the anions and the direction of fluxes remained unclear. Subsequently, a role for anion fluxes in exocytosis of neutrophil secretory compartments other than lysosomes has been hypothesized (25, 26).

The objectives of the present study were: 1) to examine Cl movements in human neutrophils exposed to particulate stimuli, 2) to investigate the potential relationships between Cl fluxes and degranulation, and 3) to characterize signal transduction pathways involved in the activation of Cl fluxes. To achieve these goals, we devised an experimental setting that may allow simultaneous assessment of both Cl fluxes and neutrophil functional responses. We provide evidence, for the first time, that phagocytosis of opsonized bacteria, fungi, and zymosan, elicits a Cl efflux similar to that induced by known soluble Cl releasers, such as TNF and fMLP. Such an efflux is due to a net outward movement of Cl, because it is accompanied by a marked, irreversible loss of Cli. We also show that Cl efflux regulates the release of azurophilic granules and the changes in cytosolic Ca2+ levels, and depends on protein tyrosine phosphorylation.

Percoll was obtained from Pharmacia Biotech. BSA, [2,3-dichloro-4-(2-methylene-butyryl)phenoxy]acetic acid (ethacrynic acid, EA), 4,4′-diisothiocyanatostilbene-2,2′-disulfonic acid (DIDS), d-glucuronic acid sodium salt, 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB), 2-[[3-(trifluoromethyl)phenyl]-amino]-3-pyridinecarboxylic acid (niflumic acid, NA), 3-p-trifluoromethylphenoxy-3-phenyl-N-methyl-propylamine hydrochloride (fluoxetine, FLX), anthracene-9-carboxylic acid (9-AC), (o-[3-hydroxymercuri-2-methoxy- propyl)carbamoyl]phenoxyacetic acid (mersalyl, MA), α-cyano-4-hydroxycinnamic acid (CHC), fMLP, cytochalasin D, pertussis toxin (Ptx), Sabouraud dextrose broth, PBS, and zymosan were obtained from Sigma-Aldrich. 1-(6-{[17β-3-methoxyoesta-1,3,5(10)-trien-17-yl]amino}hexyl)-1H-pyrrole-2,5-dione (U73122) and 1-(6-{[17β-3-methoxyoesta-1,3,5(10)-trien-17-yl]amino}hexyl)-pyrrolidine-2,5-dione (U73343) were purchased from BIOMOL. Recombinant human TNF produced in Escherichia coli was obtained from Santa Cruz Biotechnology. Fura 2-AM and 5-(and 6)-carboxyfluorescein succinimidyl ester (5(6)-FAM-SE) were purchased from Molecular Probes Europe BV. 5(6)-FAM-SE was dissolved in DMSO at 20 mg/ml, and stored as a stock solution in the dark at −20°C. High-purity trypan blue (TB) (color index 23850) was obtained from Merck KgaA. The dye was dissolved in distilled water at 5 mg/ml and filtered through a Millipore filter to remove nonsolubilized material. Na36Cl (sp. act., 14–15 μCi/g Cl) was purchased from Amersham International. All other reagents and chemicals were of the highest purity grade available. All solutions were made in LPS-free water or physiologic saline (0.9% NaCl) for clinical use.

Neutrophils were isolated from peripheral blood anticoagulated with EDTA (final concentration, 4 mM) obtained from blood donors or laboratory personnel. Isolation was made according to the method described by Metcalf et al. (27), with slight modifications. In brief, 4 ml of anticoagulated fresh blood was layered onto a discontinuous Percoll gradient consisting of 4 ml of 62% and 4 ml of 75% Percoll in PBS and centrifuged at 200 × g for 10 min and then, without interruption, at 400 × g for an additional 15 min. Neutrophils were collected at the interface between the 62 and 75% Percoll and washed once at 250 × g for 7 min with Ca2+- and Mg2+ -free HEPES-buffered saline (HBS) solution (145 mM NaCl, 5 mM KCl, 5 mM HEPES buffer, 5 mM glucose, pH 7.4). A brief (10-s) hypotonic lysis of contaminating erythrocytes was performed by suspending the cells in 3 ml of 1 mM phosphate buffer. Isotonicity was then restored by adding 7 ml of 1.3% NaCl in 1 mM phosphate buffer. After an additional washing in HBS containing 0.2% BSA (HBS-BSA), the isolated neutrophils were resuspended in the same medium at the desired concentration. The resulting cell population contained 95–97% neutrophils, 2–3% eosinophils, and 1–2% lympho-monocytes. Before starting the functional assays, CaCl2 and MgCl2 (final concentration of both cations, 1 mM) were added to the neutrophil suspension. Incubation procedures were performed in poly(2-hydroxyethyl methacrylate) (poly[HEMA]) test tubes to avoid cell sticking to the tube walls. Where indicated, neutrophils suspensions were preincubated with chloride transport blockers, or control buffer, for 15 min at 37°C.

Neutrophils, suspended at 10–15 × 106/ml in Ca2+- and Mg2+-free HBS-BSA, were loaded with 36Cl as previously described by Simchowitz and De Weer (1) by incubating the cells with 36Cl (3.0–4.0 μCi/ml) for 2 h at 37°C in a shaking water bath. After loading, the cells were washed twice with prewarmed unlabeled buffer to remove the tracer and suspended in the same medium at 2.5 × 106/ml. Measurements of 36Cl efflux were performed in cells at rest or exposed to particulate or soluble stimuli. At the desired times, 800-μl aliquots of cell suspension were collected from duplicate tubes and centrifuged for 4 min at 400 × g. Then, 750-μl aliquots of the supernatants were withdrawn, and their radioactivity was counted by liquid scintillation counting in a beta counter (LS6000TA; Beckman Instruments, Fullerton, CA). The percentage of efflux was calculated as follows: [(cpm in the supernatant of timex sample) − (cpm in the supernatant of t0 sample)]/[(total cpm of cell suspension) − (cpm in the supernatant of t0 sample)] × 100.

Changes in Cli, that reflect net Cl movements, were measured using either 36Cl (4) or a chemical colorimetric method (28). In the former case, the assay was performed exactly as previously described (8). In brief, neutrophils were suspended at 4–6 × 106/ml in HBS-BSA and incubated with 3.0–4.0 μCi/ml 36Cl for 2 h at 37°C. After loading, the cells were used without washing them free of the tracer. At the selected times of incubation, 200 μl of the cell suspensions were collected from duplicate tubes, diluted into 1400 μl of HBS-BSA prewarmed at 37°C, and centrifuged for 15 s at 12,000 × g. The pellet was suspended in 60 μl of HBS-BSA, and 50-μl aliquots were diluted into 500 μl of the same buffer layered on 600 μl of a mixture (ratio 4:1, density 1.005 ± 0.001 g/L) of silicone oil (density 1.041 g/L; silicone AR 200 fluid, 200 centistokes; Serva Electrophoresis) and paraffin oil (density: 0.873 g/L; Merck). After 1 min of centrifugation at 12,000 × g, the supernatants were discarded, the bottom of the tubes were cut, and the cell pellet-associated radioactivity was counted as described above. Background values of 36Cl trapped within the intercellular spaces in the cell pellet was measured by centrifuging an aliquot of the cell suspension immediately after the addition of Na36Cl for the loading procedure. This radioactivity was found to account for <0.5% of the total radioactivity of the cell pellet after 2 h of loading with the tracer, and therefore, it was omitted from the calculations of 36Cli. The 36Cl that remained associated to neutrophils at the selected incubation time was expressed as a percentage of 36Cl associated to neutrophils at t = 0.

Cli was determined also by means of a commercial kit (Chlorofix; Menarini Diagnostics) based on the colorimetric chemical assay described by Schoenfeld and Lowellen (28). This method was used instead of the radioactivity-based technique described above to avoid overestimation of cell-associated 36Cl caused by entrapment within the phagosome(s) of the radiotracer present in the extracellular medium. The Chlorofix chemical assay is based on the reaction of Cl with mercuric thiocyanate with generation of SCN. This anion, in turn, reacts with Fe3+ and produces a reddish-brown end product that is proportional to Cl concentration and can be read at 460 nm. Fig. 1 shows a calibration curve obtained with different concentrations of NaCl. The Chlorofix chemical assay proved to be sensitive enough to detect Cl concentrations in the range expected to be measured in our experimental conditions. Preliminary experiments showed that changes in Cli in TNF- or PMA-stimulated neutrophils as measured by using either 36Cl or the Chlorofix reagent, gave similar results, thus proving the reliability of the chemical assay (data not shown). The assay was performed as follows. At the selected incubation times, aliquots of neutrophil suspensions (5–10 × 106/ml in HBS-BSA) were collected and centrifuged for 4 min at 400 × g. The resulting pellets were suspended in 2 ml of isotonic sucrose and subsequently centrifuged for 30 s at 3500 × g to wash out extracellular Cl. The supernatants were carefully removed and discarded whereas the pellets were suspended in 100 to 200 μl of distilled water to lyse the cells and free intracellular Cl. After 20 min of incubation at room temperature, the cell lysates were subjected to a freeze-and-thaw cycle to fully disrupt the cells and disperse cell debris. The samples were centrifuged for 15 s at 12,000 × g and aliquots of the supernatants (25–75 μl) were subsequently tranferred in duplicate wells of a microtiter plate. After addition of the Chlorofix reagent (200 μl), the plate was incubated for 2–3 min at room temperature, and absorbance was then read at 460 nm in a microplate reader (Power Wave X; BioTek Instruments). Cli measured in neutrophils at the selected incubation time was expressed as a percentage of Cli at t = 0. Mean basal Cl concentration (Cli at t = 0), as assessed by Chlorofix reagent, was 82.5 ± 19.8 mM (mean ± SD, n = 5), a value in agreement with previously published data (1, 2). In the experiments with C. albicans, control assays were performed in the absence of neutrophils to determine the contribution of candida-associated Cl to the overall Cl content of the samples. Candida-associated Cl was found to account for <1.5% of the total Cl content, and therefore, it was neglected in the calculations of Cli.

FIGURE 1.

Determination of standard chloride concentrations by means of Chlorofix reagent. The assay was performed as detailed in Materials and Methods.

FIGURE 1.

Determination of standard chloride concentrations by means of Chlorofix reagent. The assay was performed as detailed in Materials and Methods.

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Staphylococcus aureus 502A (American Type Culture Collection 27217) was a generous gift of Dr. J. Iandolo (Department of Microbiology and Immunology, University of Oklahoma Health Science Center, Oklahoma City, OK). C. albicans was a clinical isolate kindly provided by Dr. E. Crevatin (Unità Clinico Operativa di Igiene e Medicina Preventiva, University of Trieste).

S. aureus was collected from single colonies grown on Luria-Bertani (LB)-agar plates, inoculated into LB broth, and grown for 18 h at 37°C. Frozen aliquots of C. albicans blastospores were diluted in Sabouraud broth and grown overnight at 30°C. Microorganisms were pelleted by centrifugation at 2000 × g for 5 min, transferred into a microtube, washed once in 0.9% NaCl solution by centrifugation at 12,000 × g for 10 s, and suspended in 1.5 ml of 0.9% NaCl. S. aureus concentration was assessed by measurments of turbidity at 500 nm. Blastospore concentrations were determined by microscopic counting in a cell-counting chamber. S. aureus and C. albicans suspensions were diluted to 1 × 108 cells/ml and 6 × 107 cells/ml, respectively, in HBS supplemented with 1 mM CaCl2 and 1 mM MgCl2 (Ca2+/Mg2+-HBS) and opsonized with 10% pooled human serum for 30 min at 37°C in a shaking water bath. Opsonized microorganisms were kept on ice until use. Where indicated, C. albicans blastospores were opsonized with human purified Ig (>95% IgG) (IG VENA; Kedrion S.p.A.) or IgG-free pooled human serum provided by Dr. L. Rigonat (Department of Physiology and Pathology, University of Trieste) and prepared by immunoabsorption on protein G-Sepharose column (Pharmacia LKB Biotechnology). Depletion of IgG from the eluted serum was ascertained by means of ELISA performed with rabbit anti-human IgG polyclonal Abs (Sigma-Aldrich) (data not shown). Zymosan was suspended in 0.9% NaCl and heated in a boiling water bath for 15 min. After centrifugation, the pellet was resuspended in Ca2+/Mg2+-HBS and opsonized with 10% pooled human serum for 30 min at 37°C. Serum-treated zymosan (STZ) was then washed with HBS, suspended in the same medium and counted electronically (Coulter Counter Mod ZBI; Coulter Electronics).

Labeling of C. albicans blastospores was performed according to the method described by Dri et al. (29), with slight modifications. Serum-opsonized yeast particles were diluted in NaHCO3 buffer (0.1 M, pH 8.3), washed and suspended in the same medium (300 × 106/ml) containing 1.5 μg/ml 5(6)-FAM-SE to start labeling. After 30 min of incubation at 4°C in the dark under continuous stirring, the reaction was stopped by adding an equal volume of 50 mM glycine in 0.9% NaCl, pH 8.5, to remove and quench weakly bound 5(6)-FAM-SE molecules. After a further 30 min of incubation at 4°C in the dark under stirring, C. albicans particles were washed twice at 2000 × g for 7 min in PBS, suspended in the same medium, and stored at −20°C.

Neutrophils suspended at 2.5 × 106/ml in Ca2+/Mg2+-HBS were incubated at 37°C in a shaking water bath with opsonized, 5(6)-FAM-SE-labeled C. albicans (1:3 neutrophil to yeast ratio). Unless otherwise stated, the mixtures were incubated for 30 min at 37°C. To stop phagocytosis, aliquots of the incubation mixtures were withdrawn into the tubes used for fluorescence flow cytometry (FCM) analysis containing an equal volume of 250 μg/ml TB dissolved in ice-cold 0.1 M citrate buffer (pH 4.0). In this way, sample pH was lowered to nearly 4.0, thereby optimizing the TB quenching effect (30). After 1 min of incubation in ice, the samples were analyzed by FCM.

FCM analysis of neutrophils incubated with fluorescence-labeled yeasts was conducted according to the method described by Busetto et al. (31). FCM was performed with a FACSCalibur (BD Biosciences), equipped with an air-cooled 15-mW argon-ion laser, operating at 488 nm. 5(6)-FAM-SE green fluorescence (FL1) was collected using a 530 ± 30 bandpass filter; red fluorescence emitted from TB bound to 5(6)-FAM-SE-labeled C. albicans particles (FL3) was collected by using a 650 ± 13 bandpass filter. The data were collected using linear amplification for forward scattering and side scattering, and logarithmic amplification for FL1 and FL3. Each sample was analyzed for 30 s at the slowest flow rate to minimize the coincidental appearance of free yeasts and neutrophils in the laser beam. The data were then analyzed by using CellQuest software from BD Biosciences. A forward scatter threshold was set to gate out debris electronically by analyzing sample buffer alone. Neutrophils and free C. albicans particles were discriminated by combined measurements of forward scattering and side scattering and gated in R1 and R2 regions, respectively. The percentage distribution of neutrophil subsets was calculated from dot plot analysis (FL1 vs FL3) of R1-gated events.

Degranulation was measured after exposure of 36Cl-loaded neutrophils to opsonized C. albicans or fMLP to simultaneously assess granule release and chloride movements. Assay conditions were as described for 36Cl efflux evaluation. In the experiments with fMLP, neutrophils were preincubated for 5 min at 37°C with cytochalasin D (2.5 μg/ml, final concentration). At the selected incubation times, aliquots of the cell suspensions were withdrawn and immediately centrifuged for 4 min at 400 × g. The supernatant fluids were collected and stored on ice. Neutrophils contained in the pellets were disrupted in 0.1% (w/v) of cetyltrimethylammonium bromide. Both supernatants and disrupted cells were transferred into duplicate wells and the activity of the selected granule markers was assayed as detailed below. Release of granular proteins was expressed as a percentage of total activity.

Colorimetric quantitative assays of enzyme activities were conducted in duplicate wells of microtiter plates. Myeloperoxidase (MPO) activity was measured as previously described (32), using 3,3′,5,5′-tetramethylbenzidine (TMB) as peroxidase substrate. In brief, 75 μl of 20 mM acetate buffer (pH 5.5) containing 2 mM TMB and 0.1% (w/v) cetyltrimethylammonium bromide were deposited into the wells. Then, 75 μl of either supernatant fluid or 20-fold diluted cell lysate were added. The peroxidatic reaction was started with H2O2 (0.3 mM final concentration) and blocked after 2–3 min with 100 μl of 2 N H2SO4. Absorbance was then read at 405 nm.

Elastase activity was assayed according to the method described by Ciesla et al. (33), by cleavage of the specific elastase substrate N-methoxysuccinyl-ala-ala-pro-val-p-nitroanilide. The incubation mixture included: 270 μl of 100 mM Tris-HCl buffer (pH 7.5) containing 10 mM CaCl2, 500 mM NaCl, 40% (v/v) DMSO, 30 μl of either supernatant fluid or diluted cell lysate, and 18 μl of N-methoxysuccinyl-ala-ala-pro-val-p-nitroanilide dissolved at 1 mg/ml in DMSO. After incubating the plate overnight at 37°C, absorbance was read 405 nm.

β-glucuronidase was assayed according to Gianetto and de Duve (34), using phenolphthalein glucuronide as substrate. Aliquots (50–100 μl) of supernatant fluid or diluted neutrophil lysates were transferred into the wells containing 90 μl of 200 mM sodium acetate buffer (pH 5.0) and 10 μl of phenolphthalein glucuronide dissolved at 10 mM in distilled water. After incubating the plate overnight at 37°C, absorbance was read at 550 nm.

Release of the specific granule marker lactoferrin (LF) was measured by ELISA using anti-human LF rabbit polyclonal Abs (Sigma-Aldrich). All steps were conducted at room temperature. Immobilization of Abs onto plate microwells with hydrazide surface (HZ wells) (carbohydrate-binding 8-well Strip Plate, catalog no. 2508; Costar), which allows site-specific binding of Abs through the carbohydrate moieties of the Fc region (35), was performed exactly as described by Menegazzi et al. (9). Supernatant fluids or cell lysates were diluted 300-fold in PBS containing 0.1% (w/v) BSA and 0.1% (v/v) Tween 20 (Merck) (PBS-T-BSA). 10 to 50 μl of the diluted samples were transferred into duplicate HZ-wells and incubated for 45 min. After 3 washings with PBS-T-BSA, biotinylated anti-human LF rabbit polyclonal Abs were added to the wells and the plate was incubated for further 45 min. After three additional washings, 50 μl of HRP-conjugated streptavidin (diluted 1000-fold in PBS-T-BSA) were added to the wells. After 45 min, HZ-wells were extensively rinsed with PBS-T-BSA and the peroxidatic reaction was subsequently conducted as described (32), using TMB as substrate. After blocking the peroxidatic reaction with 2 N H2SO4, absorbance was read at 405 nm with a microplate reader (Power Wave X; BioTek Instruments). LF concentrations in the samples were calculated using a calibration curve set up with known amounts of human LF. In our experimental conditions, LF cellular content was nearly 2.8 μg/106 cells.

Surface expression of CD18, a marker of secretory vesicles, secondary and tertiary granules (36), was measured in neutrophils incubated in suspension for 30 min at 37°C in either Ca2+/Mg2+-HBS or Ca2+/Mg2+-HBS gluconate buffer. On completion of incubation, the cell suspensions were cooled on ice for 10 min and subsequently incubated for a further 60 min with 5 μg/ml mAb TS1-18 recognizing the CD18 subunit (common β-chain) of the CD11/CD18 Ag complex (β2 integrins) (37). The cells were then washed free of the mAbs in ice-cold PBS and incubated for 45 min with a FITC-labeled rabbit anti-mouse IgG F(ab′)2 (Sigma-Aldrich). After two washings in PBS, the cells were suspended in 1% formaldehyde and analyzed by FCM using a FACSCalibur (BD Biosciences) flow cytometer.

The release of the cytoplasmic enzyme lactate dehydrogenase was assayed as an index of cytotoxicity. The assay was performed by measuring the consumption of NADH during the conversion of pyruvate to lactate as described by Bergmeyer et al. (38).

In these experiments, fura-2 AM (Molecular Probes) was used as the fluorescent, Ca2+-sensitive indicator. Neutrophils (5–10 × 106/ml in HBS buffer) were incubated in suspension with 2 μM fura-2 AM for 20 min at 37°C, diluted 5-fold and further maintained at 37°C for an additional 20 min. On completion of the loading procedure, the cells were diluted 1:1 with HBS buffer and washed twice in the same buffer. Neutrophils were suspended at 2.5 × 106/ml in HBS buffer containing 1.0 mM Ca2+ and kept in the dark at room temperature. In the experiments in which Ca2+ and 36Cl fluxes were measured in the same cell suspension, loading with fura-2 AM was conducted during the last 40 min of incubation of neutrophils with Na36Cl (see above). For each assay, 1 ml of the cell suspension was put in a thermostated cuvette in a 650-10S fluorescence spectrophotometer (PerkinElmer) equipped with a device for continuous stirring of the incubation mixture. Excitation and emission wavelengths were set at 340 and 510 nm, respectively. Before starting the assay, MgCl2 (final concentration, 1 mM) was added to the cell suspension. After a stable baseline was obtained, the cells were exposed to either opsonized C. albicans blastospores or fMLP. Increase in fura-2 fluorescence was monitored over a 5- to 10-min period. To assay the effect of Cl transport inhibition on changes in cytosolic Ca2+, neutrophils were preincubated with the selected inhibitors, or control buffer, for 15 min at 37°C in the cuvette under continuous stirring. Such a preincubation did not modify the neutrophil basal Ca2+ level (data not shown).

Neutrophils were fixed in 2% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) for 15 min at 4°C. For peroxidase staining, the cells were rinsed twice with 0.05 M Tris-HCl buffer (pH 7.5) at room temperature and were then incubated in the same buffer containing 0.05% 3,3′-diaminobenzidine tetrahydrochloride and 0.02% H2O2 (freshly prepared) for 60 min. On completion of the incubation, the cells were rinsed with 0.1 M cacodylate buffer (pH 7.4) for 10 min and postfixed in 2% OsO4 for 60–90 min at room temperature. Neutrophils were then dehydrated through a series of graded ethanols and embedded in Dow epoxy resin (DER 332; Unione Chimica Europea). Ultrathin sections were cut by an Ultrathome III (Pharmacia LKB), stained with lead citrate, and examined with a transmission electron microscope (TEM) (EM208; Philips). For quantitative evaluation, at least 150 cell sections showing nuclear lobes were scored.

Statistical significance was tested by Student’s t tests calculated using GraphPad Prism 3.0 (GraphPad Software). Values of p < 0.05 were considered statistically significant.

It is well known that human neutrophils respond to both soluble physiologic agonists (such as cytokines and chemotactic substances) and nonphysiological activators (i.e., phorbol esters, A23187) with a rapid Cl efflux (4, 5, 6, 7, 8). Because Cl movements in phagocytosing neutrophils have not yet been rigorously evaluated, we have performed a series of experiments to address this issue. Specifically, 36Cl-loaded neutrophils were exposed to three distinct serum-opsonized phagocytosable particles, namely STZ, S. aureus and C. albicans. Fig. 2,A shows that these particles triggered a similar efflux of 36Cl. Notably, this efflux was comparable to that observed in cells exposed to the soluble agonists TNF and fMLP. To establish whether phagocytosis-induced Cl efflux is accompanied by a decrease in Cli, or otherwise results from activation of Cl/Cl exchange (1), we subsequently measured variations in Cli. As shown in Fig. 2,B, Cli of resting neutrophils did not change significantly throughout the incubation. In contrast, phagocytosis caused a marked decrease of Cli, thereby indicating a net outward flux of Cl. Additionally, Fig. 2 B demonstrates that the Cl ions lost after phagocytosis were not regained. In contrast, the markedly reduced levels of Cli caused by treatment with fMLP and TNF returned to normal at 60 min after exposure.

FIGURE 2.

Serum-opsonized phagocytosable particles stimulate a net outward chloride efflux in human neutrophils. After loading with 36Cl, neutrophils were washed twice with prewarmed HBS-BSA buffer to remove the tracer and suspended at 2.5 × 106/ml in HBS-BSA buffer containing 1 mM CaCl2 and 1 mM MgCl2. A, Release of 36Cl was measured in the supernatants of cells at rest (rest) or exposed to serum-opsonized particles and soluble stimuli. Cell-to-particle ratios were 1:10, 1:3, and 1:5 for STZ, C. albicans and S. aureus, respectively. fMLP and TNF were used at the concentration of 10 nM and 0.6 nM, respectively. After 20 min of incubation at 37°C in a shaking water bath, 800-μl aliquots of cell suspensions were collected from duplicate tubes and centrifuged for 4 min at 400 × g. 750-μl aliquots of the supernatants were then withdrawn for radioactivity counting. Data represent the mean ± SD of three to eleven experiments. B, Cli was measured in neutrophils at rest or stimulated with either C. albicans (cell-to-particle ratio: 1:3), 10 nM fMLP or 0.6 nM TNF by using the Chlorofix chemical assay (for details see Materials and Methods). At the selected incubation time, aliquots of neutrophil suspensions were collected and centrifuged for 4 min at 400 × g. The resulting pellets were suspended in isotonic sucrose and subsequently centrifuged for 30 s at 3500 × g to wash out extracellular Cl. The pellets were suspended in distilled water to lyse the cells and release intracellular Cl. After 20 min of incubation at room temperature followed by a freeze-and-thaw cycle, the samples were centrifuged for 15 s at 12,000 × g and aliquots of the supernatants (25–75 μl) were subsequently transferred in duplicate wells of a microtiter plate. After addition of the Chlorofix reagent (200 μl), the plate was incubated for 2–3 min at room temperature and absorbance was read at 460 nm in a microplate reader. Cli measured in neutrophils at the selected incubation times was expressed as a percentage of Cli at t = 0. Neutrophil mean basal Cl concentration (Cli at t = 0) was 82.5 ± 19.8 mM (mean ± SD, n = 5). Data represent the mean ± SEM of three to five experiments.

FIGURE 2.

Serum-opsonized phagocytosable particles stimulate a net outward chloride efflux in human neutrophils. After loading with 36Cl, neutrophils were washed twice with prewarmed HBS-BSA buffer to remove the tracer and suspended at 2.5 × 106/ml in HBS-BSA buffer containing 1 mM CaCl2 and 1 mM MgCl2. A, Release of 36Cl was measured in the supernatants of cells at rest (rest) or exposed to serum-opsonized particles and soluble stimuli. Cell-to-particle ratios were 1:10, 1:3, and 1:5 for STZ, C. albicans and S. aureus, respectively. fMLP and TNF were used at the concentration of 10 nM and 0.6 nM, respectively. After 20 min of incubation at 37°C in a shaking water bath, 800-μl aliquots of cell suspensions were collected from duplicate tubes and centrifuged for 4 min at 400 × g. 750-μl aliquots of the supernatants were then withdrawn for radioactivity counting. Data represent the mean ± SD of three to eleven experiments. B, Cli was measured in neutrophils at rest or stimulated with either C. albicans (cell-to-particle ratio: 1:3), 10 nM fMLP or 0.6 nM TNF by using the Chlorofix chemical assay (for details see Materials and Methods). At the selected incubation time, aliquots of neutrophil suspensions were collected and centrifuged for 4 min at 400 × g. The resulting pellets were suspended in isotonic sucrose and subsequently centrifuged for 30 s at 3500 × g to wash out extracellular Cl. The pellets were suspended in distilled water to lyse the cells and release intracellular Cl. After 20 min of incubation at room temperature followed by a freeze-and-thaw cycle, the samples were centrifuged for 15 s at 12,000 × g and aliquots of the supernatants (25–75 μl) were subsequently transferred in duplicate wells of a microtiter plate. After addition of the Chlorofix reagent (200 μl), the plate was incubated for 2–3 min at room temperature and absorbance was read at 460 nm in a microplate reader. Cli measured in neutrophils at the selected incubation times was expressed as a percentage of Cli at t = 0. Neutrophil mean basal Cl concentration (Cli at t = 0) was 82.5 ± 19.8 mM (mean ± SD, n = 5). Data represent the mean ± SEM of three to five experiments.

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Opsonophagocytosis encompasses two main steps, namely 1) recognition, i.e., particle binding to phagocyte surface receptors including FcγR(s) and/or complement receptors (CR1/CR3), and 2) particle internalization requiring dynamic modifications of the actin cytoskeleton (39, 40, 41, 42, 43). To assess the relative role of binding and internalization in opsonophagocytosis-induced Cl efflux, we have conducted experiments in which binding, ingestion and Cl movements were simultaneously measured. We used C. albicans in such experiments inasmuch as we have recently developed a cytofluorometric assay to simultaneously assess its recognition and ingestion (31).

Neutrophil-candida mixtures were incubated in suspension, and after a predetermined time period, aliquots were withdrawn and assayed for 36Cl efflux, recognition and ingestion. Fig. 3,A shows that the microfilament-disrupting agent cytochalasin D did not inhibit phagocytosis-induced Cl efflux. In contrast, this agent abrogated internalization of yeasts without affecting binding to neutrophils (Fig. 3,B). Altogether these findings indicate that opsonin receptor-mediated recognition is sufficient to elicit Cl efflux even in the absence of internalization. As demonstrated in Fig. 4, serum-derived opsonins are required to induce Cl efflux by C. albicans. Accordingly, nonopsonized yeasts did not elicit Cl efflux. Additionally, opsonization with purified IgGs or IgG-depleted serum, used as a source of complement-derived opsonins, was almost as effective as opsonization with whole serum in inducing Cl efflux.

FIGURE 3.

Recognition, but not ingestion, is required to trigger 36Cl efflux in neutrophils during phagocytosis of serum-opsonized C. albicans. 36Cl-loaded neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were incubated for 5 min at 37°C in a shaking water bath without or with 2.5 μg/ml cytochalasin D. After addition of fluorescence-labeled, serum opsonized-C. albicans particles (cell-to-particle ratio, 1:3), the incubation was prolonged for additional 30 min. Aliquots of the incubation mixtures were then withdrawn and assayed for both 36Cl efflux (A) and phagocytosis (B). Assay of 36Cl efflux was as described in Fig. 2. Quantification of recognition and ingestion of candida particles by neutrophils was performed by FCM using the fluorescence quenching technique described in Materials and Methods. Data are the mean ± SEM of three to six experiments.

FIGURE 3.

Recognition, but not ingestion, is required to trigger 36Cl efflux in neutrophils during phagocytosis of serum-opsonized C. albicans. 36Cl-loaded neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were incubated for 5 min at 37°C in a shaking water bath without or with 2.5 μg/ml cytochalasin D. After addition of fluorescence-labeled, serum opsonized-C. albicans particles (cell-to-particle ratio, 1:3), the incubation was prolonged for additional 30 min. Aliquots of the incubation mixtures were then withdrawn and assayed for both 36Cl efflux (A) and phagocytosis (B). Assay of 36Cl efflux was as described in Fig. 2. Quantification of recognition and ingestion of candida particles by neutrophils was performed by FCM using the fluorescence quenching technique described in Materials and Methods. Data are the mean ± SEM of three to six experiments.

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FIGURE 4.

IgG- or C3b/bi-coated C. albicans particles, but not uncoated particles, trigger 36Cl efflux in neutrophils during phagocytosis. C. albicans blastospores were opsonized with either 10% pooled human serum (whole s), 10% IgG-free pooled human serum (IgG-depl s), 5 mg/ml purified human Ig (>95% IgG) (purified h-IgG) or incubated with medium alone (NO, nonopsonized) for 30 min at 37°C. Assay of 36Cl efflux was as in Fig. 2. Data represent the mean ± SEM of three experiments performed with neutrophils isolated from different donors.

FIGURE 4.

IgG- or C3b/bi-coated C. albicans particles, but not uncoated particles, trigger 36Cl efflux in neutrophils during phagocytosis. C. albicans blastospores were opsonized with either 10% pooled human serum (whole s), 10% IgG-free pooled human serum (IgG-depl s), 5 mg/ml purified human Ig (>95% IgG) (purified h-IgG) or incubated with medium alone (NO, nonopsonized) for 30 min at 37°C. Assay of 36Cl efflux was as in Fig. 2. Data represent the mean ± SEM of three experiments performed with neutrophils isolated from different donors.

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The regulatory role of Cl movements in several neutrophil functions is well established. Accordingly, fluctuations of Cli have been shown to promote conformational changes and to increase expression of β2 integrins, TNF- and β2 integrin-mediated spreading and metabolic activation, as well as cytoplasmic alkalinization (5, 6, 7, 8, 9). However, little is known concerning the relationships between Cl efflux and degranulation (7). We have addressed this issue in an experimental model of neutrophils phagocytosing serum-opsonized C. albicans. Under these circumstances, both degranulation (44, 45, 46) and Cl ions efflux occur. First, several structurally unrelated chloride transport blockers (1, 47, 48, 49, 50, 51, 52) were screened for their effect on particle recognition and ingestion. We performed these experiments to identify drugs that did not affect phagocytosis. Based on our results (Table I), we excluded MA and CHC inasmuch as these compounds altered phagocytosis at any concentration tested. DIDS, 9-AC, and EA could be used at concentrations not exceeding 30 μM, 1 mM, and 150 μM, respectively. In contrast, NA, NPPB, and FLX were usable at all the concentrations tested.

Table I.

Effect of Cl transport inhibitors on phagocytosis of serum-opsonized C. albicana

Cell TreatmentResponse (%)
RecognitionIngestion
Control bufferb 100.0 100.0 
EA (μM)   
 50 103.1 105.6 
 150 101.2 101.0 
 250 60.1 36.9 
DIDS (μM)   
 30 101.3 100.8 
 60 55.2 48.0 
 120 51.8 36.5 
MA (μM)   
 500 150.1 140.5 
 1000 180.6 150.0 
 2500 160.5 145.4 
CHC (μM)   
 500 130.7 140.4 
 1000 150.1 160.4 
 2500 145.8 70.6 
9-AC (μM)   
 500 101.8 100.4 
 1000 102.2 101.2 
 2500 45.7 55.2 
NA (μM)   
 50 102.5 100.1 
 100 108.0 95.9 
 200 105.1 106.2 
NPPB (μM)   
 50 99.3 101.9 
 100 93.0 103.1 
 150 97.0 103.1 
FLX (μM)   
 12.5 100.9 99.2 
 25 106.5 102.5 
 50 95.2 96.3 
Cell TreatmentResponse (%)
RecognitionIngestion
Control bufferb 100.0 100.0 
EA (μM)   
 50 103.1 105.6 
 150 101.2 101.0 
 250 60.1 36.9 
DIDS (μM)   
 30 101.3 100.8 
 60 55.2 48.0 
 120 51.8 36.5 
MA (μM)   
 500 150.1 140.5 
 1000 180.6 150.0 
 2500 160.5 145.4 
CHC (μM)   
 500 130.7 140.4 
 1000 150.1 160.4 
 2500 145.8 70.6 
9-AC (μM)   
 500 101.8 100.4 
 1000 102.2 101.2 
 2500 45.7 55.2 
NA (μM)   
 50 102.5 100.1 
 100 108.0 95.9 
 200 105.1 106.2 
NPPB (μM)   
 50 99.3 101.9 
 100 93.0 103.1 
 150 97.0 103.1 
FLX (μM)   
 12.5 100.9 99.2 
 25 106.5 102.5 
 50 95.2 96.3 
a

Neutrophils (2.5 × 106/ml) were incubated for 15 min at 37°C in a shaking water bath with the indicated substances and subsequently mixed with serum-opsonized, 5(6)-FAM-SE-labeled C. albicans (1:3 cell-to-candida ratio). After 30 min of incubation at 37°C, aliquots of the incubation mixtures were diluted with an equal volume of 250 μg/ml TB dissolved in ice-cold 0.1 M citrate buffer (pH 4) and analyzed by flow cytometry. Quantitation of recognition and ingestion were performed as described in detail in Materials and Methods. Data are the means of two independent experiments performed with neutrophils isolated from different donors.

b

Control assays were performed with cells suspended in HBS buffer containing the appropriate working dilution of the solvent. Stock solutions of all inhibitors were made in DMSO, except MA, which was dissolved in physiologic saline.

Fig. 5 shows that five of six compounds (namely EA, NA, 9-AC, NPPB, and FLX) coordinately inhibited 36Cl efflux (A) as well as the release of the azurophilic granule marker MPO (B). These compounds, however, did not significantly affect the release of the specific granule marker LF (C). DIDS did not alter 36Cl efflux or granule release at the tested concentration. None of the compounds affected lactate dehydrogenase release or inhibited the enzymatic activity of purified human MPO (data not shown). These results indicated that inhibition of MPO release could not be due to drug-induced cell toxicity or alterations in enzymatic activity of MPO. Cl transport blockers inhibited also the release of elastase and β-glucuronidase, two additional azurophilic granule markers (inset in Fig. 5 B).

FIGURE 5.

Effect of Cl transport inhibitors on 36Cl efflux and degranulation of neutrophils phagocytosing serum-opsonized C. albicans. 36Cl-loaded neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were pretreated for 15 min at 37°C in a shaking water bath with the indicated Cl transport inhibitors (EA, 150 μM; DIDS, 30 μM; NA, 200 μM; 9-AC, 1000 μM; NPPB, 150 μM; FLX, 50 μM) or with control buffer containing 0.3% DMSO. After a further 30-min incubation in the presence of serum-opsonized C. albicans (cell-to-particle ratio, 1:3), aliquots of the cell suspensions were withdrawn and assayed for 36Cl efflux (A) and release of azurophilic (B) and specific (C) granule markers. Assay of 36Cl efflux was as in Fig. 2. The release of MPO, elastase, β-glucuronidase and LF were assessed as described in Materials and Methods. Data in A, B, and C are means ± SEM of five to eleven experiments. Data in the inset of B represent the mean of two independent experiments (in duplicate) performed with cells isolated from different donors. Asterisks in A and B denote values significantly different from the control (∗∗, p < 0.01; ∗∗∗, p < 0.001).

FIGURE 5.

Effect of Cl transport inhibitors on 36Cl efflux and degranulation of neutrophils phagocytosing serum-opsonized C. albicans. 36Cl-loaded neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were pretreated for 15 min at 37°C in a shaking water bath with the indicated Cl transport inhibitors (EA, 150 μM; DIDS, 30 μM; NA, 200 μM; 9-AC, 1000 μM; NPPB, 150 μM; FLX, 50 μM) or with control buffer containing 0.3% DMSO. After a further 30-min incubation in the presence of serum-opsonized C. albicans (cell-to-particle ratio, 1:3), aliquots of the cell suspensions were withdrawn and assayed for 36Cl efflux (A) and release of azurophilic (B) and specific (C) granule markers. Assay of 36Cl efflux was as in Fig. 2. The release of MPO, elastase, β-glucuronidase and LF were assessed as described in Materials and Methods. Data in A, B, and C are means ± SEM of five to eleven experiments. Data in the inset of B represent the mean of two independent experiments (in duplicate) performed with cells isolated from different donors. Asterisks in A and B denote values significantly different from the control (∗∗, p < 0.01; ∗∗∗, p < 0.001).

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The effect of Cl transport inhibitors on neutrophil degranulation during phagocytosis was subsequently investigated at the ultrastructural level. Neutrophils pretreated with either solvent (0.3% DMSO) or 150 μM EA, a broad spectrum and potent inhibitor of Cl movements (1, 4, 6, 7, 8, 9), were incubated for 15 min with C. albicans. Preparations were then fixed, stained for peroxidase, and subsequently analyzed by TEM. The results of these analyses showed that the percentage of phagocytosing cells in EA-treated neutrophils (calculated as the number of cell sections showing at least one ingested candida particle) was similar to that observed in control neutrophils (nearly 60%). These data are in keeping with the results of cytofluorometric studies (Table I). One representative experiment is shown in Fig. 6. In >90% of control neutrophils, the peroxidase-positive granules gathered around and/or fused with the phagosome membrane (Fig. 6, ac). Conversely, azurophilic granules were scattered within the cytoplasm of EA-treated cells and only occasionally they were fused with the phagosomal membrane (Fig. 6, df). In contrast, specific granules fused with phagosomes were evident in EA-treated neutrophils (Fig. 6 f and inset therein).

FIGURE 6.

TEM analysis of neutrophils phagocytosing serum-opsonized C. albicans. Neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were treated with 0.3% DMSO (ac) or 150 μM EA (df) for 15 min at 37°C and subsequently incubated for further 15 min in the presence of C. albicans (cell-to-particle ratio, 1:3). The cell suspensions were then centrifuged for 4 min at 400 × g and the pellets were fixed in 2% (v/v) glutaraldehyde diluted in 0.1 M cacodylate buffer, pH 7.4. After three washing steps, two in cacodylate buffer and one in 0.05 M Tris-HCl buffer (pH 7.5), the cells were resuspended in the same buffer and stained for peroxidase using 3,3′-diaminobenzidine tethrahydrochloride. For additional experimental details, see Materials and Methods. For quantitative evaluation, at least 150 cell sections showing nuclear lobes were scored. Pictures obtained in one representative experiment are shown. Similar results were obtained in two other separate experiments. C, C. albicans; AG, azurophilic granules; SG, specific granules.

FIGURE 6.

TEM analysis of neutrophils phagocytosing serum-opsonized C. albicans. Neutrophils (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2) were treated with 0.3% DMSO (ac) or 150 μM EA (df) for 15 min at 37°C and subsequently incubated for further 15 min in the presence of C. albicans (cell-to-particle ratio, 1:3). The cell suspensions were then centrifuged for 4 min at 400 × g and the pellets were fixed in 2% (v/v) glutaraldehyde diluted in 0.1 M cacodylate buffer, pH 7.4. After three washing steps, two in cacodylate buffer and one in 0.05 M Tris-HCl buffer (pH 7.5), the cells were resuspended in the same buffer and stained for peroxidase using 3,3′-diaminobenzidine tethrahydrochloride. For additional experimental details, see Materials and Methods. For quantitative evaluation, at least 150 cell sections showing nuclear lobes were scored. Pictures obtained in one representative experiment are shown. Similar results were obtained in two other separate experiments. C, C. albicans; AG, azurophilic granules; SG, specific granules.

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Taken together, these results suggest a causal relationship between Cl efflux and degranulation. Moreover, they indicate that degranulation of diverse secretory compartments are differentially sensitive to changes in Cli. This hypothesis was further substantiated by data shown in Table II, obtained in cells with artificially lowered Cli concentrations. In fact, suspension of resting cells for 30 min in Cl-free buffer resulted in a nearly 40% decrease of basal Cli, as well as in different mobilization of secretory organelles. Specifically, compared with cells suspended in Cl-containing buffer, the release of secretory vesicles, tertiary and secondary granules—measured as surface expression of CD18 (36)—was markedly higher (+430%) compared with the release of LF (+187%) as well as of MPO (+98%).

Table II.

Cli decrease in resting neutrophils is associated with the release of secretory compartments

Cell TreatmentCli (percentage of basal)aCD18 ExpressionbLF release (%)cMPO release (%)d
HBS buffer 92.1 26.3 7.8 6.7 
HBS gluconate buffer 61.6 139.4 22.4 13.3 
Cell TreatmentCli (percentage of basal)aCD18 ExpressionbLF release (%)cMPO release (%)d
HBS buffer 92.1 26.3 7.8 6.7 
HBS gluconate buffer 61.6 139.4 22.4 13.3 
a

Neutrophils (5 × 106/ml in HBS-BSA) were incubated with 3 μCi/ml Na36Cl for 2 h at 37°C. To assay changes in 36Cli of neutrophils suspended in HBS buffer, the cells were used without washing. To assay changes in 36Cli of neutrophils suspended in Cl-free buffer, 36Cl-loaded cells were diluted 50 times in HBS-gluconate buffer (145 mM sodium gluconate, 5 mM potassium gluconate, 5 mM HEPES buffer, 5 mM glucose, pH 7.4), centrifuged and suspended in the same buffer. This washing step did not affect the basal 36Cli (6200 vs 6050 cpm/107 cells for washed and nonwashed cells, respectively; mean of duplicate assays). After 30 min of incubation at 37°C in a shaking water bath, 200 μl of the cell suspensions were collected from the tubes, diluted and subjected to washing procedures, as detailed in Materials and Methods. The 36Cl that remained associated to neutrophils was expressed as a percentage of 36Cl asssociated to cells at t = 0. Data are the means of two experiments (in duplicate) performed with cells isolated from different donors.

b

CD18 expression was measured by immunofluorescence flow cytometry, as described in detail in Materials and Methods. Data represent mean channel fluorescence values.

c

Assays of LF and MPO

d

release were performed as described in Fig. 5. The amount of LF and MPO released in the extracellular medium are expressed as percentage of the total activities.

Ca2+ plays a crucial role in the regulation of neutrophil degranulation. Accordingly, neutrophil secretory compartments have been shown to degranulate in hierarchical order depending on their sensitivity to cytosolic Ca2+ concentrations. In this regard, azurophilic granules display the highest Ca2+ threshold. In fact, proteins contained within these granules are released only when cytosolic Ca2+ concentrations rise to 700 nM or higher (21, 53).

Because inhibition of Cl efflux influenced the release of azurophilic but not specific granules, we asked whether such difference could be ascribed to a regulatory effect of Cl fluxes on cytosolic Ca2+ levels. To address this issue, the effects of chloride channels blockers on changes in cytosolic Ca2+ and 36Cl efflux were simultaneously assessed. In these experiments, EA and FLX—belonging to the phenoxyacetate and propylamine family, respectively—were used. Neutrophil suspensions were loaded with both 36Cl and fura-2 AM, washed, transferred into a cuvette, and then incubated either with or without EA or FLX under continuous stirring. After the addition of opsonized C. albicans, changes in cytosolic Ca2+ were recorded in a spectrophotofluorometer. An aliquot of the cell suspension was centrifuged and the resulting supernatant was assayed for 36Cl. Fig. 7, A–C shows that both EA and FLX inhibited 36Cl efflux and cytosolic Ca2+ elevation in a dose-dependent manner. Intriguingly, the calculated IC50 values for inhibition of cytosolic Ca2+ and 36Cl efflux were similar (EA, 117.1 ± 3.5 μM and 125.0 ± 1.2 μM; FLX, 31.1 ± 3.5 μM and 34.3 ± 3.1 μM for Ca2+ and 36Cl, respectively, mean ± SD, n = 3–6). As expected, both compounds inhibited azurophilic granules release by nearly 50% as determined by MPO release (data not shown).

FIGURE 7.

Effect of Cl transport inhibitors on phagocytosis-induced cytosolic Ca2+ elevation and 36Cl efflux. Neutrophils were loaded with 36Cl and fura-2 AM, washed in HBS buffer, and supended at 2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2. A total of 1 ml of the cell suspension was then transferred into a cuvette kept at 37°C, placed in a fluorescence spectrophotometer equipped with a device for continuous stirring. A, To assay the effect of EA and FLX on cytosolic Ca2+ elevation and 36Cl efflux, neutrophils were preincubated without and with the compounds, for 15 min under continuous stirring After a stable baseline was obtained, the cells were exposed to serum-opsonized C. albicans and changes in fura-2 fluorescence were monitored over a 10 min period. An aliquot of the cell suspension was withdrawn, centrifuged and the supernatant assayed for 36Cl efflux as described in Fig. 2. Data of intracellular Ca2+ and 36Cl efflux are expressed as percentage of control fura-2 fluorescence and 36Cl release, respectively. B and C, Representative fura-2 recordings of candida-induced cytosolic Ca2+ changes in neutrophils exposed to different concentrations of FLX and EA. The arrows indicate the addition of serum-opsonized candida blastospores. D, E, and F, 36Cl- and fura-2-loaded neutrophils were suspended in Ca2+-free HBS-BSA containing 1 mM MgCl2. EGTA (1 mM, final concentration) was added to the incubation mixture 2 min before the addition of candida blastospores. Changes in cytosolic Ca2+ and 36Cl efflux were monitored as above. Assay of recognition was as described in Fig. 3. Data are means ± SEM of three to ten experiments. Asterisks in E and F denote values significantly different from the control (∗, p < 0.05; ∗∗, p < 0.01).

FIGURE 7.

Effect of Cl transport inhibitors on phagocytosis-induced cytosolic Ca2+ elevation and 36Cl efflux. Neutrophils were loaded with 36Cl and fura-2 AM, washed in HBS buffer, and supended at 2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2. A total of 1 ml of the cell suspension was then transferred into a cuvette kept at 37°C, placed in a fluorescence spectrophotometer equipped with a device for continuous stirring. A, To assay the effect of EA and FLX on cytosolic Ca2+ elevation and 36Cl efflux, neutrophils were preincubated without and with the compounds, for 15 min under continuous stirring After a stable baseline was obtained, the cells were exposed to serum-opsonized C. albicans and changes in fura-2 fluorescence were monitored over a 10 min period. An aliquot of the cell suspension was withdrawn, centrifuged and the supernatant assayed for 36Cl efflux as described in Fig. 2. Data of intracellular Ca2+ and 36Cl efflux are expressed as percentage of control fura-2 fluorescence and 36Cl release, respectively. B and C, Representative fura-2 recordings of candida-induced cytosolic Ca2+ changes in neutrophils exposed to different concentrations of FLX and EA. The arrows indicate the addition of serum-opsonized candida blastospores. D, E, and F, 36Cl- and fura-2-loaded neutrophils were suspended in Ca2+-free HBS-BSA containing 1 mM MgCl2. EGTA (1 mM, final concentration) was added to the incubation mixture 2 min before the addition of candida blastospores. Changes in cytosolic Ca2+ and 36Cl efflux were monitored as above. Assay of recognition was as described in Fig. 3. Data are means ± SEM of three to ten experiments. Asterisks in E and F denote values significantly different from the control (∗, p < 0.05; ∗∗, p < 0.01).

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Ca2+ mobilization from inositol 1,4,5-triphosphate (Ins(1,4,5)P3)-sensitive intracellular stores may occur upon engagement of FcγR(s) and CR3 (54, 55) but not of CR1 (56, 57). Store-derived Ca2+ mobilization is an early event contributing to the overall Ca2+i elevation and the subsequent activation of several neutrophil responses, including phagocytosis-induced degranulation (58, 59). To investigate whether Cl efflux may play a role in the mobilization of Ca2+ from intracellular stores, neutrophils were preincubated with either EA or FLX in Ca2+-free, EGTA-containing buffer and exposed to opsonized C. albicans particles. Fig. 7 shows that inhibition of 36Cl efflux (E) was accompanied by inhibition of cytosolic Ca2+ elevation (F). It is worth noting that these experimental conditions did not influence particle recognition (D).

Studies with cells exposed to a soluble secretagogue (fMLP) further support a regulatory role of Cl fluxes in Ca2+ homeostasis. Fig. 8,A shows that four chemically unrelated Cl transport blockers inhibited fMLP-induced 36Cl efflux. Specifically, EA, NA, and NPPB showed the most potent inhibitory effect (nearly 70% inhibition). Inhibition by FLX was less marked although remaining statistically significant (nearly 30% inhibition, p < 0.025, n = 3). The effects of two Cl transport blockers, namely EA (strong inhibitor) and FLX (less potent inhibitor) on intracellular Ca2+ changes were then examined. Fig. 8 shows that, similar to the effects observed during phagocytosis of C. albicans (Fig. 7), inhibition of 36Cl efflux was paralleled by an inhibition of cytosolic Ca2+ elevation either in the presence (Fig. 8, B and C) or in the absence (Fig. 8,D) of extracellular Ca2+. The data reported in Table III indicate that both EA and FLX inhibited fMLP-induced neutrophil degranulation. As expected, MPO release was more markedly affected than LF release, because azurophilic granules are less sensitive to Ca2+ elevation than the specific ones (21). With fMLP as a stimulatory agent, higher concentrations of EA (300 μM) could be used compared with phagocytosis experiments. Under these circumstances, EA strongly inhibited both 36Cl efflux and cytosolic Ca2+ elevation (92% and 89%, respectively; means of two independent experiments). Accordingly, the release of both MPO and LF was also markedly reduced (95.7 ± 5.7% and 92.2 ± 7.4% inhibition, respectively; means ± SD, n = 4).

FIGURE 8.

Effect of Cl transport inhibitors on fMLP-induced 36Cl efflux and cytosolic Ca2+ elevation. Neutrophils were loaded with 36Cl and fura-2 AM as described in Fig. 7. A, 36Cl-loaded cells (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and MgCl2) were exposed for 15 min at 37°C in a shaking water bath to the indicated Cl transport inhibitors (EA, 150 μM; NA, 200 μM; NPPB, 150 μM; FLX, 50 μM), or control buffer containing 0.3% DMSO. fMLP (final concentration, 1 nM) was then added to stimulate the cells. After a 10-min incubation, aliquots of the cell suspensions were centrifuged, and the supernatants derived thereof were assayed for 36Cl as described in Fig. 2. Data are expressed as percentage of control 36Cl release and represent the mean ± SD of three to five experiments. B, Neutrophils loaded with fura-2 AM were washed and suspended in HBS-BSA containing 1 mM Ca2+ and 1 mM Mg2+. Aliquots of the cell suspension were transferred into a cuvette placed in a fluorescence spectrophotometer and treated for 15 min at 37°C with control buffer, 150 μM EA or 50 μM FLX under continuous stirring. After a stable baseline was obtained, the cells were stimulated with 1 nM fMLP. C, Representative fura-2 recordings of fMLP-induced cytosolic Ca2+ changes in neutrophils pretreated with FLX or EA. The arrow indicates the addition of the stimulus. D, Effect of EA and FLX on fMLP-induced cytosolic Ca2+ changes in neutrophils suspended in Ca2+-free buffer containing 1 mM EGTA. Asterisks in A denote values significantly different from the control (∗, p < 0.025; ∗∗, p < 0.01). Data in B and D are expressed as percentage of control fluorescence and are the means of two independent experiments performed with cells isolated from different donors.

FIGURE 8.

Effect of Cl transport inhibitors on fMLP-induced 36Cl efflux and cytosolic Ca2+ elevation. Neutrophils were loaded with 36Cl and fura-2 AM as described in Fig. 7. A, 36Cl-loaded cells (2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and MgCl2) were exposed for 15 min at 37°C in a shaking water bath to the indicated Cl transport inhibitors (EA, 150 μM; NA, 200 μM; NPPB, 150 μM; FLX, 50 μM), or control buffer containing 0.3% DMSO. fMLP (final concentration, 1 nM) was then added to stimulate the cells. After a 10-min incubation, aliquots of the cell suspensions were centrifuged, and the supernatants derived thereof were assayed for 36Cl as described in Fig. 2. Data are expressed as percentage of control 36Cl release and represent the mean ± SD of three to five experiments. B, Neutrophils loaded with fura-2 AM were washed and suspended in HBS-BSA containing 1 mM Ca2+ and 1 mM Mg2+. Aliquots of the cell suspension were transferred into a cuvette placed in a fluorescence spectrophotometer and treated for 15 min at 37°C with control buffer, 150 μM EA or 50 μM FLX under continuous stirring. After a stable baseline was obtained, the cells were stimulated with 1 nM fMLP. C, Representative fura-2 recordings of fMLP-induced cytosolic Ca2+ changes in neutrophils pretreated with FLX or EA. The arrow indicates the addition of the stimulus. D, Effect of EA and FLX on fMLP-induced cytosolic Ca2+ changes in neutrophils suspended in Ca2+-free buffer containing 1 mM EGTA. Asterisks in A denote values significantly different from the control (∗, p < 0.025; ∗∗, p < 0.01). Data in B and D are expressed as percentage of control fluorescence and are the means of two independent experiments performed with cells isolated from different donors.

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Table III.

Effect of Cl transport inhibitors on fMLP-induced degranulationa

Cell TreatmentMPO Release (%)LF Release (%)
Inhibition (%)Inhibition (%)
Solventb 40.6 ± 5.3 56.6 ± 6.1 
150 μM EA 8.7 ± 4.5 78.6 38.4 ± 3.2 32.1 
50 μM FLX 19.9 ± 5.6 51.0 54.3 ± 5.1 4.0 
Cell TreatmentMPO Release (%)LF Release (%)
Inhibition (%)Inhibition (%)
Solventb 40.6 ± 5.3 56.6 ± 6.1 
150 μM EA 8.7 ± 4.5 78.6 38.4 ± 3.2 32.1 
50 μM FLX 19.9 ± 5.6 51.0 54.3 ± 5.1 4.0 
a

Neutrophils (2.5 × 106/ml) were preincubated for 15 min at 37°C in a shaking water bath with the Cl transport inhibitors. Cytochalasin D (2.5 μg/ml, final concentration) was added during the last 5 min of preincubation. The cells were subsequently exposed to 1 nM fMLP for 15 min. On completion of the incubation, aliquots of the cell suspensions were withdrawn, centrifuged for 4 min at 400 × g and the supernatant fluids were collected and stored on ice. MPO and LF were assayed in supernatants and lysed pellets as detailed in Materials and Methods. Data are the mean ± SEM of three to five experiments.

b

0.3% DMSO.

These results suggest that Cl fluxes may act as an upstream regulatory signal for cytosolic Ca2+ elevation triggered by activation of different receptor types. If this is the case, it is expected that Cl efflux is independent of Ca2+ elevation. To test whether actually Cl efflux is independent, 36Cl efflux was assayed in neutrophils loaded with the Ca2+ buffering agent BAPTA and subsequently suspended in Ca2+-free medium containing EGTA. As expected (Fig. 9,A) Ca2+-depleted neutrophils did not show a rise in cytoplasmic Ca2+ after exposure to either opsonized candida particles or fMLP. Furthermore, Ca2+ depletion did not alter chloride fluxes stimulated by fMLP or by phagocytosis of IgG- or C3b/bi-coated candida blastospores (Fig. 9 B). The small decrease in chloride efflux after phagocytosis of candida is likely to be due to the slight reduction in binding (∼15%) of opsonized particles to BAPTA/EGTA-treated cells (data not shown). Altogether these findings provide strong evidence that Cl efflux in stimulated neutrophils is largely independent of cytosolic Ca2+ changes.

FIGURE 9.

36Cl efflux triggered by particulate or soluble stimuli is independent of cytosolic Ca2+ elevation. After loading with 36Cl and fura-2, neutrophils were washed and suspended at 2.5 × 106/ml in HBS-BSA containing 1 mM MgCl2. The cell suspension was then divided in two aliquots, which were treated with 10 μM BAPTA or control buffer for 30 min at 37°C in a shaking water bath. On completion of the incubation, the two cell populations were centrifuged and the pellets were suspended in either HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2 (Ca2+ buffer) or HBS-BSA containing 1 mM MgCl2 and 1 mM EGTA (BAPTA/EGTA). Neutrophils were exposed for 10 min to control buffer (rest), IgG- (C-IgG), C3b/bi- (C-C3b/bi)-coated or uncoated (NO, non opsonized) candida blastospores (cell to particle ratio: 1:3), or stimulated with 1 nM fMLP. A, Changes in cytosolic Ca2+ and B, efflux of 36Cl, were assayed as described in Fig. 7. Results represent the means ± SD of three to four experiments.

FIGURE 9.

36Cl efflux triggered by particulate or soluble stimuli is independent of cytosolic Ca2+ elevation. After loading with 36Cl and fura-2, neutrophils were washed and suspended at 2.5 × 106/ml in HBS-BSA containing 1 mM MgCl2. The cell suspension was then divided in two aliquots, which were treated with 10 μM BAPTA or control buffer for 30 min at 37°C in a shaking water bath. On completion of the incubation, the two cell populations were centrifuged and the pellets were suspended in either HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2 (Ca2+ buffer) or HBS-BSA containing 1 mM MgCl2 and 1 mM EGTA (BAPTA/EGTA). Neutrophils were exposed for 10 min to control buffer (rest), IgG- (C-IgG), C3b/bi- (C-C3b/bi)-coated or uncoated (NO, non opsonized) candida blastospores (cell to particle ratio: 1:3), or stimulated with 1 nM fMLP. A, Changes in cytosolic Ca2+ and B, efflux of 36Cl, were assayed as described in Fig. 7. Results represent the means ± SD of three to four experiments.

Close modal

As previously shown, Cl ions movements triggered by particulate or soluble stimuli appear to precede and to be independent of cytosolic Ca2+ increase, one of the earliest events in the signaling pathways elicited by FcγR(s), CR3, and fMLP receptor (fMLP-R) (55, 60, 61, 62, 63, 64). We thus sought to investigate the signals involved in the control Cl fluxes. Specifically, we focused our attention on known critical intracellular messengers that act upstream of calcium, including tyrosine kinases (TKs), PI3K, PLC, and the pertussis toxin (Ptx)-sensitive Gi protein (55, 60, 61, 62, 63, 64, 65, 66, 67, 68, 69, 70, 71). To determine whether these regulatory proteins could play a role in the activation of Cl movements, 36Cl-loaded neutrophils were pretreated with specific inhibitors and subsequently exposed to opsonized candida blastospores or fMLP. Experimental results are summarized in Fig. 10.

FIGURE 10.

Activation of 36Cl efflux from neutrophils exposed to opsonized candida blastospores and fMLP: effect of inhibitors of distinct signal transduction pathways. After loading with 36Cl, neutrophils were washed and suspended at 2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2. Aliquots of the cell suspension were then treated for 15 min at 37°C in a shaking water bath with either 50 μM genistein (gen), 25 nM wortmannin (wtm), or 2.5 μM U73122 (U731). Neutrophil treatment with 1 μM Ptx was conducted for 3 h at 37°C during cell loading with 36Cl. Neutrophils were subsequently exposed to IgG- (C-IgG), C3b/bi- (C-C3b/bi) coated or uncoated (NO, nonopsonized) candida blastospores (cell-to-particle ratio: 1:3) (top panel), or 1 nM fMLP (bottom panel). Results represent the means ± SD of three experiments.

FIGURE 10.

Activation of 36Cl efflux from neutrophils exposed to opsonized candida blastospores and fMLP: effect of inhibitors of distinct signal transduction pathways. After loading with 36Cl, neutrophils were washed and suspended at 2.5 × 106/ml in HBS-BSA containing 1 mM CaCl2 and 1 mM MgCl2. Aliquots of the cell suspension were then treated for 15 min at 37°C in a shaking water bath with either 50 μM genistein (gen), 25 nM wortmannin (wtm), or 2.5 μM U73122 (U731). Neutrophil treatment with 1 μM Ptx was conducted for 3 h at 37°C during cell loading with 36Cl. Neutrophils were subsequently exposed to IgG- (C-IgG), C3b/bi- (C-C3b/bi) coated or uncoated (NO, nonopsonized) candida blastospores (cell-to-particle ratio: 1:3) (top panel), or 1 nM fMLP (bottom panel). Results represent the means ± SD of three experiments.

Close modal

Fig. 10 shows that the broad-spectrum protein TK inhibitor genistein almost completely abolished 36Cl efflux triggered by either IgG- or C3b/bi-coated C. albicans (top panel), thereby suggesting that activation of cytosolic TK is necessary for such an efflux to occur. It is worth noting that genistein, at the concentration used in these experiments, did not influence recognition and ingestion of candida particles as judged by cytofluorometric analysis (76.5 and 73.9% of cells with bound/ingested candida particles in untreated and genistein-treated cells, respectively; mean of two duplicate experiments). In contrast, fMLP-mediated Cl efflux was insensitive to genistein (bottom panel), thereby indicating that Cl channel/transporter activation is independent of the tyrosine phosphorylation wave elicited by this compound (72 and references therein).

PI3K has been shown to act downstream of cytosolic TK(s) and fMLP-R-coupled Gi protein but upstream of Ca2+ release from intracellular stores (61, 62, 63, 64, 68). It is thus possible that this kinase may act as an activator of Cl efflux. As shown in Fig. 10, wortmannin, a specific inhibitor of PI3K when used in the nanomolar range (73), did not modify 36Cl efflux from neutrophils exposed to opsonized candida (top panel) or fMLP (bottom panel). As expected, wortmannin markedly inhibited Ca2+ elevation induced by engagement of either FcγR(s) or CR3 (94.3 and 80% inhibition, respectively; mean of two experiments performed with cells isolated from different donors), thereby confirming its effectiveness as a PI3K inhibitor. It is therefore concluded that this kinase is not involved in the activation of Cl movements.

Engagement of opsonin receptors and fMLP-R results in activation of PLCγ2 and PLCβ2, respectively (55, 69, 70). Both PLC isoforms catalyze the breakdown of PtdIns(4,5)P2 to generate Ins(1,4,5)P3, and diacylglycerol (DAG) (74). Besides their role in classical signal transduction, it has been recently suggested that phosphoinositides and DAG may play a role in the regulation of ion channels (75, 76, 77). Interestingly, recent data have indicated that inflammatory mediators may trigger a volume-regulated PLC-activated chloride current through a DAG-dependent Ins(1,4,5)P3-independent pathway, without the involvement of PKC-dependent phosphorylation (78). To assess the possible role of PLC in phagocytosis- and fMLP-stimulated Cl efflux, we tested the effect of the PLC inhibitor U73122, a membrane-permeable aminosteroid that interferes with other PLC-mediated neutrophil responses (55, 67, 79). Fig. 10 shows that 36Cl fluxes stimulated by opsonized candida blastospores (top panel) or fMLP (bottom panel) were insensitive to U73122, thereby suggesting that PLC activation is not required for the occurrence of Cl efflux. It is worth noting that, under the same experimental conditions, U73122 efficiently inhibited PLC-dependent cellular responses. In fact, this compound markedly reduced phagocytosis- and fMLP-induced increase of cytosolic Ca2+ (85.3 ± 5.2% and 88.0 ± 6.1% of inhibition for candida- and fMLP-mediated response, respectively; mean ± SD, n = 3). As expected, the inactive analog U73343 had no effect on both 36Cl efflux and cytosolic Ca2+ elevation (data not shown).

fMLP-R is a member of the G protein-coupled receptor superfamily of seven transmembrane spanning receptors. Upon ligand binding to fMLP-R, the heterotrimeric Gi protein dissociates into a GTP-bound α-subunit (Gα-GTP) and a βγ-subunit (reviewed in Ref. 68). Several reports have clearly demonstrated that Ptx ribosylates Gα, thereby preventing Gi activation. Accordingly, Ptx blocks several fMLP-stimulated neutrophil functions (64, 80, 81). In keeping with these data, we have shown that Ptx markedly inhibited 36Cl efflux (bottom panel). This result suggests that Gi protein activation plays a pivotal role in fMLP-induced Cl movements.

The present study is the first to examine Cl movements in human neutrophils during phagocytosis. Specifically, we have shown that phagocytosis is accompanied by a massive and irreversible decrease in Cli and that such a decrease regulates degranulation.

Cl movements in both resting and stimulated neutrophils have been previously investigated in a host of different experimental models, including cells in suspension (1, 4, 5, 6, 7, 8), cells adherent to surfaces coated with extracellular matrix proteins (8), and cells laid onto surface-bound mAbs recognizing either the common β-chain (CD18) or the individual α-chains (CD11a, CD11b, CD11c) of β2 integrins (9). Altogether, these studies have provided novel insights into the role of Cl movements in neutrophil pathophysiology. In addition, they have clearly demonstrated a positive relationship between modifications of Cli and activation of several neutrophil responses (5, 6, 7, 8, 9). Under these circumstances, we undertook this study to shed light on the role Cl movements during neutrophil phagocytosis.

In this regard, we have first shown that phagocytosis of serum-opsonized particles is accompanied by a considerable Cl efflux which is paralleled by a marked loss of Cli (Fig. 2). Differently from the effects observed with the use of soluble agonists, the decrease in Cli induced by phagocytosis appeared irreversible, thereby suggesting that the mechanisms responsible for Cl regain in phagocytosing neutrophils are unable to counteract the loss of chloride ions. In this regard, it is worth noting that Cl movements with similar features have been reported in neutrophils adherent to fibronectin-coated surfaces (8). The mechanism of Cl reuptake remains to be elucidated, and the question as to whether this mechanism does not occur in phagocytosing cells deserves further investigations.

We then aimed to gain a better understanding of Cl fluxes in phagocytosing neutrophils. We first attempted to elucidate the relationships between Cl release and phagocytosis. To address this issue, 36Cl-loaded cells incubated with serum-opsonized, fluorescein-labeled C. albicans blastospores were used. This allowed us to evaluate simultaneously both responses. Our results showed that Cl fluxes were largely independent of particle ingestion because cytochalasin D, that almost abolished particle internalization, only slightly affected Cl efflux (Fig. 3). It is thus posited that recognition of opsonized particles plays a crucial role in triggering Cl fluxes. Moreover, nonopsonized candida particles failed to stimulate chloride efflux, thereby suggesting that engagement of opsonin receptors is essential to elicit this effect. Additionally, these results ruled out a major role of other receptors involved in neutrophil-candida interactions, such as β-glucan receptors (Dectin 1) or mannose receptors (82, 83) in the stimulation of Cl efflux. Cl fluxes induced by candida particles coated with either IgG or C3b/bi alone were lower compared with those triggered by candida opsonized with whole serum (Fig. 4). This small but reproducible difference could be ascribed to the lack of FcγR(s)/CR3 cooperation in the induction of neutrophil activation (84), because particles opsonized with whole serum, IgG or C3b/bi bound similarly to neutrophils (data not shown). These results are in agreement with our previously published data showing that CR3 (CD11b/CD18) is capable of triggering Cl efflux in human neutrophils (9). In addition, the finding that IgG-opsonized candida particles stimulate Cl fluxes suggested a novel signaling property for FcγR(s).

Release of granular proteins during phagocytosis plays a role in both microbicidal activity and tissue damage. A role for permeant anions in the regulation of neutrophil degranulation has been previously hypothesized based on inhibition of lysosomal enzyme secretion by anion channel blockers (23, 24). To gain new insights into the role of Cl movements in neutrophil degranulation, we devised an experimental set-up that allowed simultaneous evaluation of both Cl fluxes and exocytosis. Experiments with the use of several chemically unrelated Cl transport inhibitors unequivocally showed the existence of a positive correlation between Cl efflux and the release of azurophilic granules (Fig. 5, A and B). Interestingly, the release of specific granules appeared largely independent of Cl fluxes, inasmuch as none of the inhibitors significantly affected LF release (Fig. 5,C). Further evidence for a distinct sensitivity of the two major types of granules to cytoplasmic Cl levels came from a different experimental approach, namely ultrastructural analysis of phagocytosis-induced degranulation. The TEM micrographs (Fig. 6) showed that fusion of azurophilic granules with phagosomes is markedly diminished in the presence of the Cl transport inhibitor EA. In contrast, fusion of specific granules was not affected, thereby confirming enzyme assays data (Fig. 5, B and C). These findings, along with the observation that chloride transport inhibitors did not affect C. albicans ingestion (Table I), seem to exclude that the reduction of azurophilic granule release might be ascribed to drug toxicity.

Altogether these results suggest that the extent of release of distinct secretory compartments may depend on Cli levels. This possibility is supported by the observation that degranulation of all secretory organelles occurred when Cli was lowered to nearly 60% of basal levels by means of incubation in Cl-free buffer (see Table II). It should be noted, however, that the release of small secretory vesicles, tertiary and secondary granules largely exceeded that of azurophilic granules. These data are in keeping with previously published observations on the spontaneous release of specific granules from resting neutrophils suspended in Cl-free gluconate buffer (85).

Although the release of primary and secondary granules may follow distinct signaling pathways (86, 87, 88, 89, 90, 91), elevation of cytosolic Ca2+ is largely recognized as a crucial step in neutrophil degranulation. Additionally, the order of release of neutrophil secretory organelles is tightly controlled by the gradual increase of cytosolic calcium (21, 22). Because Cl fluxes appeared to be involved in the regulation of phagocytosis-induced granule secretion, we addressed the question as to whether a relationship exists between Cl and Ca2+ movements. To explore this possibility, we performed a series of cross-inhibition experiments using a model allowing simultaneous assessment of 36Cl fluxes and cytosolic Ca2+ changes. We found that Cl transport blockers had parallel, dose-dependent inhibitory effects on Cl efflux and Ca2+ elevation (Fig. 7, A–C). In contrast, depletion of intracellular calcium by BAPTA/EGTA abolished Ca2+ response (Fig. 9,A) but did not significantly influence phagocytosis-induced 36Cl efflux (Fig. 9,B). Similar results were obtained when neutrophils were exposed to the soluble agonist fMLP (Figs. 8 and 9), thereby supporting a role of Cl ions fluxes in the regulation of neutrophil calcium homeostasis. Our present findings are consistent with previous data showing that Cl movements may influence Ca2+ mobilization (92, 93).

In the present report we have also identified the intracellular signaling pathways regulating Cl fluxes. As demonstrated by the experiments with genistein and Ptx (Fig. 10), Cl fluxes triggered by engagement of immune receptors and fMLP-R depend on TK and Gi protein activation, respectively. With regard to the role of TK as regulators of Cl movements, our data are in line with previous findings showing that the Src family protein TK p56lck plays a crucial role in the activation of Cl channels in human lymphocytes (94, 95). The results obtained with the use of Ptx indicate, for the first time, that Gi protein activation is required to elicit Cl fluxes evoked by fMLP. Because other signal transducers downstream of Gi protein (TK, PI3K, PLC) do not appear to be involved in fMLP-induced Cl efflux (Fig. 10, bottom panel), it remains to be established whether Gi-derived subunits may serve as direct activators of ion channels in neutrophils, as it has been previously demonstrated for the βγ subunit in other cell types (96 and references therein).

Previous data have shown that PI3K is rapidly activated by TK following FcγR(s) or CR3 engagement (61, 62, 63). Moreover, it has been also suggested to play a role in fMLP-mediated neutrophil responses (64, 68). Working from these assumptions, we sought to determine the role of this kinase in eliciting Cl movements. Wortmannin, an inhibitor of PI3K, did not affect 36Cl efflux (Fig. 10). This result strongly suggests that PI3K is not involved in the modulation of Cl fluxes.

Neutrophils express two isoforms of PtdIns(4,5)P2-specific PLC, namely PLCγ2—which is activated by tyrosine phosphorylation in response to FcγR(s) and CR3 engagement (55, 70)—and PLCβ2, whose activation depends on the Ptx-sensitive release of βγ subunit after engagement of Gi protein-coupled fMLP-R (69). Recent reports have shown that phosphoinositides and DAG may play a role in the regulation of ion channels, including a volume-sensitive PLC-activated chloride channel evoked by inflammatory mediators through a DAG-dependent Ins(1,4,5)P3-independent pathway (75, 76, 77, 78). We therefore aimed to investigate the potential role of PLC in the modulation of Cl movements. In this study we have demonstrated that 36Cl effluxes are insensitive to the PLC inhibitor U73122 (Fig. 10), thereby suggesting that the PLC/DAG system is unlikely to play a major role in the modulation of neutrophil chloride movements.

Our current report supports the concept that Cl efflux may be considered one of the earliest events in the process of neutrophil activation (4). Of interest is also the observation that rapid activation of Cl efflux occurs in other mammalian cells. Accordingly, a very quick (t1/2 < 10 s) and dramatic (nearly 50%) loss of Cli due to rapid efflux has been described in rat parotid acinar cells upon stimulation with a muscarinic cholinergic agonist (97). It has been also shown that progesterone-mediated tyrosine phosphorylation of the sperm γ-aminobutyric acidA (GABAA)-like receptor/Cl channel triggers a rapid Cl efflux regulating Ca2+ elevation during the acrosome reaction (93).

At present, it remains to be established how changes in Cli can modulate neutrophil Ca2+ homeostasis and degranulation. Because we have demonstrated that inhibitors of Cl efflux interfere with Ca2+ release from intracellular stores (Fig. 7 D), it is posited that cytoplasmic chloride levels may influence the interaction of Ins(1,4,5)P3 with its receptor located on store membranes. The observation that the binding of Ins(1,4,5)P3 to its receptor may be competitively inhibited by heparin and other polyanionic polymers (98, 99, 100) is in keeping with our hypothesis. Alternatively, changes in Cli could be involved in the activation of PLCγ2 and PLCβ2—the PLC isoforms expressed in granulocytes (55, 69, 70, 101), which in turn mediate Ins(1,4,5)P3 formation and the increase in cytosolic calcium. In this regard, a role for chloride ion as an allosteric regulator of several different proteins has been already demonstrated (102, 103, 104, 105).

A growing body of evidence supports the notion that Cl movements are involved in the regulation of neutrophil responses (5, 6, 7, 8, 9). However, several issues still remain to be elucidated. Accordingly, the mechanisms whereby Cl transport occurs across the plasma membrane remain unclear. Several chloride transporters have been described in neutrophils, including the electroneutral Cl/HCO3exchanger (1) as well as a variety of channels, such as calcium-activated channels, voltage-dependent channels, cell swelling-activated channels, and glycine-gated channels (106, 107, 108, 109). Inhibition studies may be of aid to investigate the mechanisms involved in the regulation of Cl movements. Unfortunately, many, if not all, common chloride transport inhibitors have a broad-spectrum activity (47, 51, 106, 110, 111, 112). This phenomenon limits the potential usefulness of these compounds to shed more light on the precise molecular mechanisms involved in this process. Recently published data obtained in neutrophils isolated from mice lacking ClC-3 (Clcn3(−/−) PMN) suggest a role for this channel in the regulation of neutrophil respiratory burst and phagocytosis (112). Identification of the mechanisms responsible for phagocytosis-induced Cl efflux was beyond the aims of this study. Nevertheless, the presence of chloride fluxes in Ca2+-depleted neutrophils seems to suggest that calcium-activated Cl conductance does not play a major role in this process. Although these data conflict with previous reports (5, 91), it should be noted that our findings are consistent with other studies showing that Cl efflux triggered by the cross-linking of β2 integrins occurs in a Ca2+-independent manner (9).

Another issue regarding Cl movements in neutrophils concerns chloride localization within the cell. Recently, Segal and coworkers have provided preliminary evidence to suggest that this anion is contained at high concentrations within cytoplasmic granules to be transferred thereafter into phagocytic vacuoles (113). If this is the case, it is expected that inhibition of degranulation should be accompanied by reduction in Cl fluxes. However, there are at least two observations that argue against this possibility. First, we have demonstrated that 36Cl efflux occurs normally in Ca2+-depleted cells (Fig. 9) despite a marked inhibition of degranulation (Ref. 22 and our unpublished data). Second, cytochalasin B, known to increase fMLP- and IL-8-induced degranulation, did not modify Cl efflux induced by these agonists (Ref. 4 and our unpublished results). Further investigations are mandatory to shed more light on the intracellular distribution of chloride ion in neutrophils.

Cation/anion trafficking across biological membranes is a complex phenomenon. Moreover, the role of ion movements in neutrophil activation remains a matter of debate (reviewed in Refs. 113 and 114). The present study may shed more light on the role of permeant anions, particularly chloride, in phagocyte activation.

We thank Dr. Francesca Vita and Dr. Maria Rosa Soranzo (Department of Physiology and Pathology, University of Trieste) and Claudio Gamboz (Centro Coordinamento e Sviluppo Progetti e Apparecchiature, University of Trieste) who helped us perform the transmission electron microscopy studies. This work was dedicated to the memory of Pierluigi Patriarca.

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.

2

Abbreviations used in this paper: Cli, intracellular chloride content; EA, [2,3-dichloro-4-(2-methylene-butyryl)phenoxy]acetic acid (ethacrynic acid); DIDS, 4,4′-diisothiocyanatostilbene-2,2′-disulfonic acid; NPPB, 5-nitro-2-(3-phenylpropylamino)benzoic acid; NA, 2-[[3-(trifluoromethyl)phenyl]-amino]-3-pyridinecarboxylic acid (niflumic acid); FLX, 3-p-trifluoromethylphenoxy-3-phenyl-N-methyl-propylamine hydrochloride (fluoxetine); 9-AC, anthracene-9-carboxylic acid; MA, (o-[3-hydroxymercuri-2-methoxy-propyl)carbamoyl] phenoxyacetic acid (mersalyl); CHC, α-cyano-4-hydroxy-cinnamic acid; TB, trypan blue; HBS-BSA, HEPES-buffered saline containing 0.2% BSA; STZ, serum-treated zymosan; 5(6)-FAM-SE, 5-(and 6)-carboxyfluorescein succinimidyl ester; FCM, fluorescence flow cytometry; MPO, myeloperoxidase; LF, lactoferrin; HZ, hydrazide; TEM, transmission electron microscopy; TK, tyrosine kinase; Ptx, pertussis toxin; U73122, 1-(6-{[17β-3-methoxyoesta-1,3,5(10)-trien-17-yl]amino}hexyl)-1H-pyrrole-2,5-dione; U73343, 1-(6-{[17β-3-methoxyoesta-1,3,5(10)-trien-17-yl]amino}hexyl)-pyrrolidine-2,5-dione; PLC, phospholipase C; DAG, diacylglycerol.

1
Simchowitz, L., P. De Weer.
1986
. Chloride movements in human neutrophils: diffusion, exchange, and active transport.
J. Gen. Physiol.
88
:
167
-194.
2
Ince, C., B. Thio, B. van Duijn, J. T. van Dissel, D. L. Ypey, P. C. Leijh.
1987
. Intracellular K+, Na+ and Cl concentrations and membrane potential in human monocytes.
Biochim. Biophys. Acta
905
:
195
-204.
3
Baron, D. N., S. A. Ahmed.
1969
. Intracellular concentrations of water, and of the principal electrolytes determined by the analysis of isolated human leukocytes.
Clin. Sci.
37
:
205
-219.
4
Shimizu, Y., R. H. Daniels, M. A. Elmore, M. J. Finnen, M. E. Hill, J. M. Lackie.
1993
. Agonist-stimulated Cl efflux from human neutrophils: a common phenomenon during neutrophil activation.
Biochem. Pharmacol.
45
:
1743
-1751.
5
Myers, J. B., H. F. Cantiello, J. H. Schwartz, A. I. Tauber.
1990
. Phorbol ester-stimulated human neutrophil membrane depolarization is dependent on Ca2+-regulated Cl efflux.
Am. J. Physiol.
259
:
C531
-C540.
6
Menegazzi, R., S. Busetto, R. Cramer, P. Dri, P. Patriarca.
2000
. Role of intracellular chloride in the reversible activation of neutrophil β2 integrins: a lesson from TNF stimulation.
J. Immunol.
165
:
4606
-4614.
7
Perez-Cornejo, P., J. Arreola, F-Y. Law, J. B. Schultz, P. A. Knauf.
2004
. Volume-sensitive chloride channels do not mediate activation-induced chloride efflux in human neutrophils.
J. Immunol.
172
:
6988
-6993.
8
Menegazzi, R., S. Busetto, P. Dri, R. Cramer, P. Patriarca.
1996
. Chloride ion efflux regulates adherence, spreading, and respiratory burst of neutrophils stimulated by tumor necrosis factor-α (TNF) on biologic surfaces.
J. Cell Biol.
135
:
511
-522.
9
Menegazzi, R., S. Busetto, E. Decleva, R. Cramer, P. Dri, P. Patriarca.
1999
. Triggering of chloride ion efflux from human neutrophils as a novel function of leukocyte β2 integrins: relationship with spreading and activation of the respiratory burst.
J. Immunol.
162
:
423
-434.
10
Berton, G..
1999
. Degranulation. J. I. Gallin, and R. Snyderman, eds.
Inflammation: Basic Principles and Clinical Correlates
3rd ed.
703
-719. Lippincott Williams & Wilkins, Philadelphia.
11
Faurschou, M., N. Borregaard.
2003
. Neutrophil granules and secretory vesicles in inflammation.
Microbes Infect.
5
:
1317
-1327.
12
Bainton, D. F..
1973
. Sequential degranulation of the two types of polynorphonyclear leukocyte granules during phagocytosis of microorganisms.
J. Cell Biol.
58
:
249
-264.
13
Henson, P. M., Z. G. Oades.
1975
. Stimulation of human neutrophils by soluble and insoluble immunoglobulin aggregates. Secretion of granule constituents and increased oxidation of glucose.
J. Clin. Invest.
56
:
1053
-1061.
14
Naccache, P. H., T. F. Molski, P. Borgeat, R. I. Sha’afi.
1985
. Intracellular calcium redistribution and its relationship to fMet-Leu-Phe, leukotriene B4, and phorbol ester induced rabbit neutrophil degranulation.
J. Cell Physiol.
122
:
273
-280.
15
Richter, J., T. Andersson, I. Olsson.
1989
. Effect of tumor necrosis factor and granulocyte/macrophage colony stimulating factor on neutrophil degranulation.
J. Immunol.
142
:
3199
-3205.
16
Boros, P., J. A. Odin, T. Muryoi, S. K. Masur, C. Ba, J. C. Unkeless.
1991
. IgM anti-Fc γ R autoantibodies trigger neutrophil degranulation.
J. Exp. Med.
173
:
1473
-1482.
17
Borregaard, N., L. J. Miller, T. A. Springer.
1987
. Chemoattractant-regulated mobilization of a novel intracellular compartment in human neutrophils.
Science
237
:
1204
-1206.
18
Calafat, J., T. W. Kuijpers, H. Janssen, N. Borregaard, A. J. Verhoeven, D. Roos.
1993
. Evidence for small intracellular vesicles in human blood phagocytes containing cytochrome b558 and the adhesion molecule CD11b/CD18.
Blood
81
:
3122
-3129.
19
Kjeldsen, L., H. Sengelov, K. Lollike, M. H. Nielsen, N. Borregaard.
1994
. Isolation and characterization of gelatinase granules from human neutrophils.
Blood
83
:
1640
-1649.
20
Richter, J., J. Ng-Sikorski, I. Olsson, T. Andersson.
1990
. Tumor necrosis factor-induced degranulation in adherent human neutrophils is dependent on CD11b/CD18 integrin-triggered oscillations of cytosolic Ca2+.
Proc. Nat. Acad. Sci. USA
87
:
9472
-9476.
21
Lew, D. P., A. Monod, F. A. Waldfogel, B. Dewald, M. Baggiolini, T. Pozzan.
1986
. Quantitative analysis of the cytosolic free calcium dependency of exocytosis from three subcellular compartments in intact human neutrophils.
J. Cell Biol.
102
:
2197
-2204.
22
Sengeløv, H., L. Kjeldsen, N. Borregaard.
1993
. Control of exocytosis in early neutrophil activation.
J. Immunol.
150
:
1535
-1543.
23
Korchak, H. M., B. A. Eisenstat, S. T. Hoffstein, P. B. Dunham, G. Weissmann.
1980
. Anion channel blockers inhibit lysosomal enzyme secretion from human neutrophils without affecting generation of superoxide anion.
Proc. Natl. Acad. Sci. USA
77
:
2721
-2725.
24
Korchak, H. M., B. A. Eisenstat, J. E. Smolen, L. E. Rutherford, P. B. Dunham, G. Weissmann.
1982
. Stimulus-response coupling in the human neutrophil. The role of anion fluxes in degranulation.
J. Biol. Chem.
257
:
6916
-6922.
25
Vedder, N. B., J. M. Harlan.
1988
. Increased surface expression of CD11b/CD18 (Mac-1) is not required for stimulated neutrophil adherence to cultured endothelium.
J. Clin. Invest.
81
:
676
-682.
26
Smith, R. J., B. J. Bowman, S. C. Speziale.
1986
. Interleukin-1 stimulates granule exocytosis from human neutrophils.
Int. J. Immunopharmacol.
8
:
33
-40.
27
Metcalf, J. A., J. I. Gallin, W. M. Nauseef, R. K. Root.
1986
. Preparation of cells and materials for functional assays.
Laboratory Manual of Neutrophil Function
5
Raven Press, New York.
28
Schoenfeld, R. G., C. S. Lowellen.
1964
. A colorimetric assay for determination of serum chloride.
Clin. Chem.
10
:
533
-539.
29
Dri, P., G. Presani, S. Perticarari, L. Alberi, M. Prodan, E. Decleva.
2002
. Measurement of phagosomal pH of normal and CGD-like human neutrophils by dual florescence flow cytometry.
Cytometry
48
:
159
-166.
30
Sahlin, S., J. Hed, I. Rundquist.
1983
. Differentiation between attached and ingested immune complexes by a fluorescence quenching cytofluorometric assay.
J. Immunol. Methods
60
:
115
-124.
31
Busetto, S., E. Trevisan, P. Patriarca, R. Menegazzi.
2004
. A single-step, sensitive flow cytofluorometric assay for the simultaneous assessment of membrane-bound and ingested Candida albicans in phagocytosing neutrophils.
Cytometry
58A
:
201
-206.
32
Menegazzi, R., G. Zabucchi, A. Knowles, R. Cramer, P. Patriarca.
1992
. A new, one-step assay on whole cell suspensions for peroxidase secretion by human neutrophils and eosinophils.
J. Leukocyte Biol.
52
:
619
-624.
33
Ciesla, D.J., E. E. Moore, W. L. Biffl, R. J. Gonzalez, C. C. Silliman.
2001
. Hypertonic saline attenuation of the neutrophil cytotoxic response is reversed upon restoration of normotonicity and reestablished by repeated hypertonic challenge.
Surgery
129
:
567
-575.
34
Gianetto, R., C. de Duve.
1955
. Tissue fractionation studies, 4: comparative study of the binding of acid phosphatase, β-glucuronidase and cathepsin by rat liver particles.
Biochem. J.
59
:
433
-438.
35
Brillhart, K. L., T. T. Ngo.
1991
. Use of microwell plates carrying hydrazide groups to enhance antibody immobilization in enzyme immunoassays.
J. Immunol. Methods
144
:
19
-25.
36
Lacal, P., R. Pulidogn, F. Sanchez-Madrid, F. Mollinedo.
1988
. Intracellular location of T200 and Mol glycoproteins in human neutrophils.
J. Biol. Chem.
263
:
9946
-9951.
37
Sanchez-Madrid, F., A. M. Krenky, C. F. Ware, E. Robbins, J. L. Strominger, S. Burakoff, T. S. Springer.
1982
. Three distinct antigens associated with human T-lymphocyte-mediated cytolysis: LFA-1, LGA-2, and LFA-3.
Proc. Natl. Acad. Sci. USA
79
:
7489
-7493.
38
Bergmeyer, H. U., H. Bernt, B. Hess.
1965
. Lactic dehydrogenase. H. U. Bergmeyer, ed.
Methods in Enzymatic Analysis
736
Academic Press, New York.
39
Greenberg, S., J. El Khoury, F. Di Virgilio, E. M. Kaplan, S. C. Silverstein.
1991
. Ca2+-independent F-actin assembly and disassembly during Fc receptor-mediated phagocytosis in mouse macrophages.
J. Cell Biol.
113
:
757
-767.
40
Sheterline, P., J. E. Rickard, R. C. Richards.
1984
. Fc receptor-directed phagocytic stimuli induced transient actin assembly at an early stage of phagocytosis in neutrophil leukocytes.
Eur. J. Cell Biol.
34
:
80
-87.
41
Oliver, J. M., R. D. Berlin.
1982
. Cytoskeleton-membrane interaction and the remodeling of the cell surface during phagocytosis and chemotaxis.
Adv. Exp. Med. Biol.
155
:
113
-131.
42
Howard, T. H., R. G. Watts.
1994
. Actin polymerization and leukocyte function.
Curr. Opin. Hematol.
1
:
61
-68.
43
Serrander, L., P. Skarman, B. Rasmussen, W. Witke, D. P. Lew, K. H. Krause, O. Stendhal, O. Nusse.
2000
. Selective inhibition of IgG-mediated phagocytosis in gelsolin-deficient murine neutrophils.
J. Immunol.
165
:
2451
-2457.
44
Breton-Gorius, J., D. Houssay, J. L. Vilde, B. Dreyfus.
1975
. Partial myeloperoxidase deficiency in a case of preleukaemia, II: defects of degranulation and abnormal bactericidal activity of blood neutrophils.
Br. J. Haematol.
30
:
279
-288.
45
Ferrante, A..
1989
. Tumor necrosis factor α potentiates neutrophil antimicrobial activity: increased fungicidal activity against Torulopsis glabrata and Candida albicans and associated increase in oxygen radical production and lysosomal enzyme release.
Infect. Immun.
57
:
2115
-2122.
46
Badolato, R., J. M. Wang, S. L. Stornello, A. N. Ponzi, M. Duse, T. Musso.
2000
. Serum amyloid A is an activator of PMN antimicrobial functions: induction of degranulation, phagocytosis, and enhancement of anti-Candida activity.
J. Leukocyte Biol.
67
:
381
-386.
47
Jianq, B., N. Attori, B. Liu, Y. Nakayama, K. Kitagawa, C. Inagaki.
2004
. Suppression of cell proliferation with induction of p21 by Cl channel blokers in human leukemic cells.
Eur. J. Pharmacol.
488
:
27
-34.
48
Randriamampita, C., A. Trautmann.
1987
. Ionic channels in murine macrophages.
J. Cell Biol.
105
:
761
-769.
49
Maertens, C., L. Wei, T. Voets, G. Droogmans, B. Nilius.
1999
. Block by fluoxetine of volume-regulated anion channels.
Br. J. Pharmacol.
126
:
508
-514.
50
Bretag, A. H., S. R. Dawe, D. I. Kerr, A. G. Moskwa.
1980
. Myotonia as a side effect of diuretic action.
Br. J. Pharmacol.
71
:
467
-471.
51
Parkerson, K. A., H. Sontheimer.
2003
. Contribution of chloride channels to volume regulation of cortical astrocytes.
Am. J. Physiol.
284
:
C1460
-C1467.
52
Wangemann, P., M. Wittner, A. Di Stefano, H. C. Englert, H. J. Lang, E. Schlatter, R. Greger.
1986
. Cl channel blockers in the thick ascending limb of the loop of Henle. Structure activity relationship.
Pflügers Arch.
407
:
S128
-S141.
53
Niessen, H. W. M., A. J. Verhoeven.
1992
. Differential up-regulation of specific and azurophilic granule membrane markers in electropermeabilized neutrophils.
Cell. Signal.
4
:
501
-509.
54
Fallmann, M., D. P. Lew, O. Stendhal, T. Andersson.
1989
. Receptor-mediated phagocytosis in human neutrophils is associated with increased formation of inositol phosphates and diacylglycerol: elevation in cytosolic free calcium and formation of inositol phosphates can be dissociated from accumulation of diacylglycerol.
J. Clin. Invest.
84
:
886
-891.
55
Hellberg, C., L. Molony, L. Zheng, T. Andersson.
1996
. Ca2+ signalling mechanisms of the β2 integrin on neutrophils: involvement of phospholipase Cγ2 and Ins(1,4,5)P3.
Biochem. J.
317
:
403
-409.
56
Delibrias, C. C., E. Fisher, G. Bismuth, M. D. Kazatchkine.
1992
. Expression, molecular association, and functions of C3 complement receptors CR1 (CD35) and CR2 (CD21) on the human T cell line HPB-ALL.
J. Immunol.
149
:
768
-774.
57
Ernst, J., J. L. Rosales, S. Zimmerli.
1995
. Calcium signalling initiated by CR1 (CD35) crosslinking is mediated by phagocyte receptors in cis.
Biochem. Biophys. Res. Comm.
209
:
1032
-1038.
58
Herlin, T., N. Borregaard.
1983
. Early changes in cyclic AMP and calcium efflux during phagocytosis by neutrophils from normals and patients with chronic granulomatous disease.
Immunology
48
:
17
-26.
59
Tapper, H., W. Furuya, S. Grinstein.
2002
. Localized exocytosis of primary (lysosomal) granules during phagocytosis: Role of Ca2+-dependent tyrosine phosphorylation and microtubules.
J. Immunol.
168
:
5287
-5296.
60
Löfgren, R., L. Serrander, M. Forsberg, A. Wilsson, A. Wasteson, O. Stendhal.
1999
. CR3, FcγRIIA and FcγRIIIB induce activation of the respiratory burst in human neutrophils: the role of intracellular Ca2+, phospholipase D and tyrosine phosphorylation.
Biochem. Biophys. Acta
1452
:
46
-59.
61
Vossebeld, P. J. M., C. H. E. Homburg, R. C. Schweizer, I. Ibarrola, J. Kessler, L. Koenderman, D. Roos, A. J. Verhoeven.
1997
. Tyrosine phosphorylation-dependent activation of phosphatidylinosite 3-kinase occurs upstream of Ca2+-signalling induced by Fcγ receptor cross-linking in human neutrophils.
Biochem. J.
323
:
87
-94.
62
Dewitt, S., M. B. Hallet.
2002
. Cytosolic free Ca2+ changes and calpain activation are required for β integrin-accelerated phagocytosis by human neutrophils.
J. Cell Biol.
159
:
181
-189.
63
Li, B., D. J. Allendorf, R. Hansen, J. Marroquin, C. Ding, D. E. Cramer, J. Yan.
2006
. Yeast β-glucan amplifies phagocyte killing of iC3b-opsonized tumor cells via complement receptor 3-Syk-phosphatidylinositol 3-kinase pathway.
J. Immunol.
177
:
1661
-1669.
64
Ahmed, M. U., K. Hazeki, O. Hazeki, T. Katada, M. Ui.
1995
. Cyclic AMP-increasing agents interfere with chemoattractant-induced respiratory burst in neutrophils as a result of the inhibition of phosphatidylinositol 3-kinase rather than receptor-operated Ca2+ influx.
J. Biol. Chem.
270
:
23816
-23822.
65
Kiefer, F., J. Brummell, N. Al-Alawi, S. Latour, A. Cheng, A. Veillette, S. Grinstein, T. Pawson.
1998
. The Syk protein tyrosine kinase is essential for Fcγ rceptor signaling in macrophages and neutrophils.
Mol. Cell. Biol.
18
:
4209
-4220.
66
Raeder, E. M., P. J. Mansfield, V. Hinkovska-Galcheva, J. A. Shayman, L. A. Boxer.
1999
. Syk activation initiates downstream signaling during human polymorphonuclear leukocyte phagocytosis.
J. Immunol.
163
:
6785
-6793.
67
Zheng, L., A. Sjolander, J. Eckerdal, T. Andersson.
1996
. Antibody-induced engagement of β 2 integrins on adherent human neutrophils triggers activation of p21ras through tyrosine phosphorylation of the protooncegene product Vav.
Proc. Natl. Acad. Sci. USA
93
:
8431
-8436.
68
Bokoch, G. M..
1995
. Chemoattractant signaling and leukocyte activation.
Blood
86
:
1649
-1660.
69
Jiang, H., Y. Kuang, Y. Wu, W. Xie, M. I. Simon, D. Wu.
1997
. Roles of phospholipase C β2 in chemoattractant-elicited responses.
Proc. Natl. Acad. Sci. USA
94
:
7971
-7975.
70
Dusi, S., M. Donini, V. Della Bianca, F. Rossi.
1994
. Tyrosine phosphorylation of phospholipase C-γ 2 is involved in the activation of phosphoinositide hydrolysis by Fc receptors in human neutrophils.
Biochem. Biophys. Res. Commun.
201
:
1100
-1108.
71
Lad, P. M., C. V. Olson, P. A. Smiley.
1985
. Association of the N-formyl-Met-Leu-Phe receptor in human neutrophils with a GTP-binding protein sensitive to pertussis toxin.
Proc. Natl. Acad. Sci. USA
82
:
869
-873.
72
Mócsai, A., Z. Jacus, T. Vantus, G. Berton, C. A. Lowell, E. Ligeti.
2000
. Kinase pathways in chemoattractant-induced degranulation of neutrophils: the role of p38 mitogen-activated protein kinase activated by Src family kinases.
J. Immunol.
164
:
4321
-4331.
73
Thelen, M., M. P. Wyman, H. Langen.
1994
. Wortmannin binds specifically to 1-phosphatidylinositol 3-kinase while inhibiting guanine nucleotide-binding protein-coupled receptor signaling in neutrophil leukocytes.
Proc. Natl. Acad. Sci. USA
91
:
4960
-4964.
74
Berridge, M. J., R. F. Irvine.
1989
. Inositol phosphates and cell signalling.
Nature
341
:
197
-205.
75
Venkatachalam, K., F. Zheng, D. L. Gill.
2003
. Regulation of canonical transient potential (TRPC) channel function by diacylglycerol and protein kinase C.
J. Biol. Chem.
278
:
29031
-29040.
76
Suh, B-C., T. Inoue, T. Meyer, B. Hille.
2006
. Rapid chemically induced changes of PtdIns(4,5)P2 gate KCNQ ion channels.
Science
314
:
1454
-1457.
77
Pochynyuk, O., Q. Tonq, A. Staruschenko, H. P. Ma, J. D. Stockland.
2006
. Regulation of the epithelial Na+ channel (EnaC) by phosphatidylinositides.
Am. J. Physiol.
290
:
F949
-F957.
78
Zholos, A., B. Beck, V. Sydorenko, L. Lemmonier, P. Bordat, N. Prevarskaya, R. Skryma.
2005
. Ca2+- and volume-sensitive chloride currents are differentially regulated by agonists and store-operated Ca2+ entry.
J. Gen. Physiol.
125
:
197
-211.
79
Smith, R. J., L. M. Sam, J. M. Justen, G. L. Bundy, G. A. Bala, J. E. Bleasdale.
1990
. Receptor-coupled signal transduction in human polymorphonuclear neutrophils: effects of a novel inhibitor of phospholipase C-dependent processes on cell responsiveness.
J. Pharmacol. Exp. Ther.
253
:
688
-697.
80
Krause, K. H., W. Schlegel, C. B. Wollheim, T. Andersson, F. A. Waldfogel, P. D. Lew.
1985
. Chemotactic peptide activation of human neutrophils and HL-60 cells. Pertussis toxin reveals correlation between inositol trisphosphate generation, calcium transients, and cellular activation.
J. Clin. Invest.
76
:
1348
-1354.
81
Cassatella, M. A., F. Bazzoni, M. Ceska, I. Ferro, M. Baggiolini, G. Berton.
1992
. IL-8 production by himan polymorphonuclear leukocytes: the chemoattractant formyl-methionyl-phenylalanine induces the gene expression and release of IL-8 through a pertussis toxine-sensitive pathway.
J. Immunol.
148
:
3216
-3220.
82
Danley, D. L., A. E. Hilger.
1981
. Stimulation of oxidative metabolism in murine polymorphonuclear leukocytes by unopsonized fungal cells: evidence for a mannose-specific mechanism.
J. Immunol.
127
:
551
-556.
83
Marodi, L., H. M. Korchak, R. B. Johnston, Jr.
1991
. Mechanisms of host defense against Candida species, I: phagocytosis by monocytes and monocyte-derived macrophages.
J. Immunol.
146
:
2783
-2789.
84
Zhou, M. J., E. J. Brown.
1994
. CR3 (mac-1, α M β 2, CD11b/CD18) and Fc γ RIII cooperate in generation of a neutrophil respiratory burst: requirement of Fc γ RII and tyrosine phosphorylation.
J. Cell Biol.
125
:
1407
-1416.
85
Fittschen, C., P. M. Henson.
1994
. Linkage of azurophilic granule secretion in neutrophil to chloride ion transport and endosomal transcytosis.
J. Clin. Invest.
93
:
247
-255.
86
Wanq-Iverson, P., K. B. Pryzwansky, J. K. Spitznagel, M. H. Cooney.
1974
. Bactericidal capacity of phorbol myristate acetate-treated human polymorphonuclear leukocytes.
Infect. Immun.
22
:
945
-955.
87
Tzeng, D. Y., T. F. Deuel, J. S. Huanq, S. M. Senior, L. A. Boxer, R. L. Baehner.
1984
. Platelet-derived growth factor promotes polymorphonuclear leukocyte activation.
Blood
64
:
1123
-1128.
88
Tapper, H..
1996
. The secretion of preformed granules by macrophages and neutrophils.
J. Leukocyte Biol.
59
:
613
-622.
89
Welch, H., I. Maridonneau-Parini.
1997
. Hck is activated by opsonized zymosan and A23187 in distinct subcellular fractions of human neutrophils.
J. Biol. Chem.
272
:
102
-109.
90
Abdel-Latif, D., M. Steward, D. L. Macdonald, G. A. Francis, M. C. Dinauer, P. Lacy.
2004
. Rac2 is critical for neutrophil primary granule exocytosis.
Blood
104
:
832
-839.
91
Mollinedo, F., J. Calafat, H. Janssen, B. Martin-Martin, S. M. Canchado, J. Nabokina, C. Gajate.
2006
. Combinatorial SNARE complexes modulate the secretion of cytoplasmic granules in human neutrophils.
J. Immunol.
177
:
2831
-2841.
92
Phipps, D. J., D. R. Branch, L. C. Schlichter.
1996
. Chloride-channel block inhibits T lymphocyte activation and signalling.
Cell. Signal.
8
:
141
-149.
93
Meizel, S., K. O. Turner, R. Nuccitelli.
1997
. Progesterone triggers a wave of increased free calcium during the human sperm acrosome reaction.
Dev. Biol.
182
:
67
-75.
94
Lepple-Wienhues, A., I. Szabo, T. Laun, N. K. Kaba, E. Gulbins, F. Lang.
1998
. The tyrosine kinase p56lck mediates activation of swelling-induced chloride channels in lymphocytes.
J. Cell Biol.
141
:
281
-286.
95
Szabo, I., A. Lepple-Wienhues, K. N. Kaba, M. Zoratti, E. Gulbins, F. Lang.
1998
. Tyrosine kinase-dependent activation of a chloride channel in CD95-induced apoptosis in T lymphocytes.
Proc. Nat. Acad. Sci. USA
95
:
6169
-6174.
96
Hamm, H. E..
1998
. The many faces of G protein signaling.
J. Biol. Chem.
273
:
669
-672.
97
Melvin, J. E., M. Kawaguchi, B. J. Baum, R. J. Turner.
1987
. A muscarinic agonist-stimulated chloride efflux pathway is associated with fluid secretion in rat parotid acinar cells.
Biochem. Biophys. Res. Commun.
145
:
754
-759.
98
O’Rourke, F., M. B. Feinstein.
1990
. The inositol 1,4,5-triphosphate receptor binding sites of platelet membranes. pH dependency, inhibition by polymeric sulphates, and the possible presence of arginine at the binding site.
Biochem. J.
267
:
297
-302.
99
Tanabe, T., H. Otani, L. Bao, Y. Mikami, T. Yasukura, T. Ninomiya, R. Ogawa, C. Inagaki.
1996
. Intracellular signaling pathway of substance P-induced superoxide production in human neutrophils.
Eur. J. Pharmacol.
299
:
187
-195.
100
Davies-Cox, E. V., I. Laffafian, M. B. Hallet.
2001
. Control of Ca2+ influx in human neutrophils by inositol 1,4,5-triphosphate (IP3) binding: differential effects of micro-injected IP3 receptor antagonist.
Biochem. J.
355
:
139
-143.
101
Bianchini, L., G. Todderud, S. Grinstein.
1993
. Cytosolic [Ca2+] homeostasis and tyrosine phosphorylation of phospholipase C γ 2 in HL60 granulocytes.
J. Biol. Chem.
268
:
3357
-3363.
102
Na, K. J., H. J. Lee.
1983
. Role of chloride ion as an allosteric activator of angiotensin-converting enzyme.
Arch. Biochem. Biophys.
227
:
580
-586.
103
Rohm, K. H..
1985
. Chloride as allosteric effector of yeast aminopeptidase I.
Arch. Biochem. Biophys.
239
:
216
-225.
104
Higashijima, T., K. M. Ferguson, P. C. Sternweis.
1987
. Regulation of hormone-sensitive GTP-dependent regulatory proteins by chloride.
J. Biol. Chem.
262
:
3597
-3602.
105
Cigic, B., R. H. Pain.
1999
. Location of the binding site for chloride ion activation of cathepsin C.
Eur. J. Biochem.
264
:
944
-951.
106
Schumann, M. A., P. Gardner, T. A. Raffin.
1993
. Recombinant human tumor necrosis factor α induces calcium oscillation and calcium-activated chloride current in human neutrophils.
J. Biol. Chem.
268
:
2134
-2140.
107
Stoddard, J. S., J. H. Steinbach, L. Simchowitz.
1993
. Whole cell Cl currents in human neutrophils induced by cell swelling.
Am. J. Physiol.
265
:
C156
-C165.
108
Schumann, M. A., T. A. Raffin.
1994
. Activation of a voltage-dependent chloride current in human neutrophils by phorbol 12-myristate 13-acetate and formyl-methionyl-leucyl-phenylalanine.
J. Biol. Chem.
269
:
2389
-2398.
109
Wheeler, M., R. F. Stachlewitz, S. Yamashima, K. Ikejima, A. L. Morrow, R. G. Thurman.
2000
. Glycine-gated chloride channels in neutrophils attenuate calcium influx and superoxide production.
FASEB J.
14
:
476
-484.
110
Day, Y., J. Zhang.
2004
. Chloride efflux is involved in ET-1 and 5-HT-induced contraction of rabbit basilar artery.
J. Cardiovasc. Pharmacol.
44
:
S125
-S128.
111
Huang, L., J. Cao, H. Wang, L. A. Vo, J. G. Brand.
2005
. Identification and functional characterization of a voltage-gated chloride channel and its novel splice variant in taste bud cells.
J. Biol. Chem.
280
:
36150
-36157.
112
Moreland, J. G., A. P. Davies, G. Bailey, W. M. Nauseef, F. S. Lamb.
2006
. Anion channels, including ClC-3, are required for normal neutrophil oxidative function, phagocytosis, and transendothelial migration.
J. Biol. Chem.
281
:
12277
-12288.
113
Segal, A. W..
2005
. How neutrophils kill microbes.
Annu. Rev. Immunol.
23
:
197
-223.
114
Murphy, R., T. E. DeCoursey.
2006
. Charge compensation during the phagocyte respiratory burst.
Biochem. Biophys. Acta
1757
:
996
-1011.