VSOP/Hv1 proton channels sustain calcium entry, neutrophil migration, and superoxide production by limiting cell depolarization and acidification

Voltage-activated proton currents (I_H+) were recorded in neutrophils purified from peripheral blood using acidic pipette solutions to activate I_H+ (pHi/o = 6.1/7.2). Large voltage-activated time-dependent outward currents were observed in neutrophils from WT mice (Fig. 1 A). These currents were reversibly blocked by 100 µM Zn²⁺, a known inhibitor of proton currents (unpublished data). In contrast, no detectable I_H+ was recorded from VSOP/Hv1^−/− neutrophils (Fig. 1 B). The current–voltage relationship of control neutrophils was outwardly rectifying with a threshold of activation around −30 mV, whereas the I-V curve of VSOP/Hv1^−/− neutrophils was completely flat. We and others previously suggested that the phagocyte NADPH oxidase contains a built-in proton channel (Henderson et al., 1987) that opens only when the oxidase is active (Bánfi et al., 1999; Petheo et al., 2006). Whether the oxidase is a proton channel or modulates another channel has been much debated (Maturana et al., 2002; Morgan et al., 2002; Touret and Grinstein, 2002). To settle this question, we recorded proton currents in VSOP/Hv1^−/− neutrophils activated by PMA to activate the oxidase and reveal its putative proton channel. As expected, PMA increased the amplitude of I_H+ in control neutrophils by ∼2.5-fold, accelerated its activation kinetics by the same extent, and slowed its deactivation kinetics by ∼1.7-fold (Fig. 1, D and F). In contrast, no current was recorded from VSOP/Hv1^−/− neutrophils activated with PMA (Fig. 1, E and F).

Proton channels are potent acid extruders that participate in the extrusion of the cytosolic acid generated by the phagocyte NADPH oxidase (Demaurex et al., 1996). To test whether the VSOP/Hv1 proton channel is required for acid extrusion, we measured cytosolic pH changes with BCECF during activation of neutrophils with PMA, using solutions devoid of Na⁺ and of bicarbonate ions to minimize the contribution of the Na⁺/H⁺ antiporter and Cl⁻/HCO₃ exchanger. As shown in Fig. 2, VSOP/Hv1^−/− neutrophils were slightly more acidic than control neutrophils before stimulation (pH_cyt 6.87 ± 0.13 vs. 7.13 ± 0.13, P > 0.05) and acidified markedly upon stimulation with PMA, with their cytosolic pH decreasing by 0.4 pH U, whereas control cells alkalinized slightly (6.46 ± 0.17 vs. 7.18 ± 0.15, P < 0.01; Fig. 2, B and C). These data indicate that VSOP/Hv1 proton channels extrude the acid generated in the cytosol during the activation of neutrophils by PMA.

The failure to see proton currents in neutrophils activated with PMA could reflect a failure of the patch-clamped cells to mount a functional oxidase at their plasma membrane. To ensure that this was not the case, we recorded the electron currents (Ie⁻) generated by the electrogenic activity of the enzyme (Schrenzel et al., 1998). The perforated patch configuration was used to avoid rundown of the current that could occur in whole cell mode. As shown in Fig. 3 A, addition of 0.1 µM PMA evoked within 1 min an inward current at −60 mV that was fully inhibited by the subsequent addition of 1 µM of the oxidase inhibitor diphenyliodonium (DPI), thus fulfilling the criteria for Ie⁻. Importantly, the amplitude of Ie⁻ recorded from VSOP/Hv1^−/− and WT neutrophils was identical (−2.1 ± 0.4 vs. −2.3 ± 0.7 pA, n = 11 and 6, respectively; Fig. 3 B) and approached the value of −2.6 ± 0.5 pA recorded in PMA-activated mouse granulocytes by Morgan et al. (2007). The normal Ie⁻ amplitude in VSOP/Hv1^−/− neutrophils indicates that electrons are transferred from cytosolic NADPH to extracellular oxygen at normal rates, implying that VSOP/Hv1^−/− neutrophils have a fully functional oxidase. The absence of proton currents in these cells thus proves that the oxidase does not contain a proton channel.

The normal Ie⁻ amplitude of VSOP/Hv1^−/− neutrophils appears at odds with the reduced superoxide production of these cells that was recently reported (Okochi et al., 2009; Ramsey et al., 2009). To confirm these studies, we measured the capacity of VSOP/Hv1^−/− neutrophils to produce H₂O₂. As shown in Fig. 3 C, H₂O₂ production was reduced by 74% in our VSOP/Hv1^−/− neutrophils purified from peripheral blood. Reassuringly, 1 mM of the proton channel inhibitor Zn²⁺ inhibited H₂O₂ production by control neutrophils by 80% but had only minor effects on VSOP/Hv1^−/− neutrophils. To confirm that the reduced H₂O₂ production of VSOP/Hv1^−/− neutrophils was caused by the ablation of endogenous proton channels, we exposed cells to gramicidin, a molecule that generates pores permeable to protons and other monovalent ions. As shown in Fig. 3 (C and D), gramicidin almost restored H₂O₂ production by VSOP/Hv1^−/− neutrophils to the levels of control cells. Thus, an artificial proton conductance can restore high-level H₂O₂ production in cells lacking endogenous proton channels. Activation of the oxidase depolarizes the plasma membrane to voltages as high as +60 mV in physiological saline (Jankowski and Grinstein, 1999). This depolarization is mitigated by proton channels that equilibrate the membrane potential to the H⁺ equilibrium potential (Bánfi et al., 1999). Thus, the most likely explanation for the reduced superoxide production of VSOP/Hv1^−/− neutrophils is an increased depolarization in cells lacking the charge-compensating proton channel. To test this possibility, we monitored membrane potential changes with DiBAC(4)3 during activation of neutrophils. As shown in Fig. 3 E, VSOP/Hv1^−/− neutrophils depolarized significantly more than control neutrophils after PMA stimulation. Addition of Zn²⁺ potentiated the depolarization of control but not of VSOP/Hv1^−/− neutrophils (Fig. 3 F). Subsequent addition of gramicidin collapsed the membrane potential of both control and VSOP/Hv1^−/− neutrophils to the same level (unpublished data). This indicates that the reduced H₂O₂ production of VSOP/Hv1^−/− neutrophils can be explained by the increased depolarization conferred by the lack of the proton channel.

Besides decreasing superoxide production, the increased depolarization reduces the driving force for the entry of cations into cells. The lack of VSOP/Hv1 thus likely impacts on the influx of Ca²⁺, an ion which controls numerous cellular functions. In neutrophils, Ca²⁺ entry occurs predominantly across store-operated Ca²⁺ channels (SOCEs) activated by the depletion of intracellular Ca²⁺ stores (Scharff and Foder, 1993). To test whether VSOP/Hv1 ablation could have an impact on SOCE, we measured the changes in cytosolic Ca²⁺ concentration with fura-2. Neutrophils were exposed to PMA for 15 min to maximally activate the oxidase and depolarize the plasma membrane, thapsigargin was added in the absence of Ca²⁺ to deplete intracellular calcium stores, and Ca²⁺ was then readmitted to evoke SOCE. As shown in Fig. 4 A, thapsigargin evoked similar responses in control and VSOP/Hv1^−/− neutrophils, indicating that Ca²⁺ release from stores is not altered by the loss of VSOP/Hv1 proton channels. However, SOCE was strongly reduced in VSOP/Hv1^−/− neutrophils, as expected from the higher membrane voltage of these cells (Fig. 4, A and D). To see whether the blunted SOCE could alter Ca²⁺ responses to physiological agonists, we exposed cells to the bacterial peptide N-formyl-Met-Ile-Val-Ile-Leu (fMIVIL), a potent agonist of the mouse formyl peptide receptors (Southgate et al., 2008). In the absence of preactivation with PMA, fMIVIL evoked biphasic Ca²⁺ elevations, reflecting an initial component of Ca²⁺ release followed by a delayed component of Ca²⁺ influx, as in human neutrophils (Demaurex et al., 1992). The integrated Ca²⁺ responses were significantly reduced in VSOP/Hv1^−/− neutrophils exposed to fMIVIL for 30 min (Fig. 4, B and D), mainly because of a reduction in the delayed influx component of the Ca²⁺ signal. Interestingly, the fMIVIL Ca²⁺ responses were monophasic in neutrophils preactivated with PMA, suggesting that Ca²⁺ influx activates more rapidly in neutrophils exposed to the phorbol ester, possibly because of the phosphorylation of Ca²⁺ handling proteins. Remarkably, the Ca²⁺ responses were almost abrogated in VSOP/Hv1^−/− neutrophils preactivated with PMA (Fig. 4, C and D). This indicates that VSOP/Hv1^−/− neutrophils cannot mount effective Ca²⁺ signals when their oxidase is active, most likely as a result of the decreased SOCE imparted by the enhanced depolarization.

Among other functions, Ca²⁺ ions regulate the activity of gelsolin to depolymerize the actin cytoskeleton (Larson et al., 2005). Rapid disassembly of the actin cytoskeleton is critical in neutrophils, highly motile cells which must crawl out of blood vessels to seek bacteria inside tissues. To test whether the Ca²⁺ defect of VSOP/Hv1^−/− neutrophils could impair actin depolymerization, we labeled cortical actin with fluorescent phalloidin during phagocytosis of zymosan particles. As shown in Fig. S1, a thicker ring of polymerized actin was observed around particles ingested by VSOP/Hv1^−/− neutrophils. This indicates that cortical actin depolymerization is indeed impaired in these cells, most likely as a result of their abortive calcium signals. To test whether impaired actin depolymerization could alter the mobility of VSOP/Hv1^−/− neutrophils, we measured the spreading and migration of neutrophils by transmission imaging. As shown in Fig. 5 (A and B), the majority of VSOP/Hv1^−/− neutrophils (72%) spread spontaneously on BSA, a nonadhesive substrate. In contrast, only 35% of control neutrophils spread spontaneously on BSA. We next investigated the chemokinetic movement of neutrophils induced by the addition of fMIVIL. Addition of fMIVIL caused neutrophils to move in nondirectional fashion (Videos 1 and 2) and revealed a clear migration defect in neutrophils lacking VSOP/Hv1. When exposed to fMIVIL, these cells moved more slowly and migrated a smaller distance. Tracking the migration paths of cells revealed that the averaged mobility of VSOP/Hv1^−/− neutrophils was reduced by 43%, from 4.67 ± 0.27 to 2.66 ± 0.30 µm/min (Fig. 5, C and D). Importantly, the spreading and mobility defects were observed in the absence of PMA and in cells exposed to fMIVIL alone. To test whether the oxidase was active in these conditions, we measured the production of ROS evoked by fMIVIL. As shown in Fig. S2, fMIVIL induced a transient production of H₂O₂, indicating that fMIVIL activates the oxidase, albeit less efficiently than PMA. Addition of PMA induced virtually all control and VSOP/Hv1^−/− neutrophils to spread (Fig. 5 A, bottom) and prevented their migration in response to fMIVIL. Interestingly, VSOP/Hv1^−/− neutrophils often exhibited a long tail that appeared stuck to the plate (Video 2), suggesting that their slower mobility was the result of an inability to detach from the substrate. To prove the causal relationship between the calcium and migration defects, we added the calcium ionophore ionomycin during fMIVIL-induced neutrophil migration, using a low dose of ionomycin (100 nM) and a low extracellular Ca²⁺ concentration (∼5 µM) to minimize cytotoxicity. As shown in Fig. 5 D, addition of ionomycin slightly improved the mobility of neutrophils from WT mice, confirming that the cytosolic Ca²⁺ clamp was effective and was not cytotoxic. Strikingly, the migration defect of VSOP/Hv1^−/− neutrophils was fully restored by the addition of ionomycin (Fig. 5 D). These data indicate that lack of VSOP/Hv1 proton channels impedes the migration of neutrophils by aborting physiological Ca²⁺ signals.

Our findings establish the mechanism by which VSOP/Hv1 proton channels sustain the activity of the phagocyte NADPH oxidase and reveal that these channels control other neutrophil functions. Cells lacking the VSOP/Hv1 molecule exhibited normal electron currents when stimulated with PMA and produced normal levels of H₂O₂ when an artificial proton conductance was provided by gramicidin. This indicates that these cells contain a fully functional oxidase able to transfer electrons efficiently across the plasma membrane. Yet VSOP/Hv1^−/− neutrophils failed to produce adequate amounts of H₂O₂ when their membrane voltage was not clamped with a patch-clamp electrode or with an ionophore. We generated two pieces of evidence that can account for the reduced H₂O₂ production in VSOP/Hv1^−/− neutrophils stimulated with PMA: (1) The lack of VSOP/Hv1 was associated with a substantial acidification of the cytosol, demonstrating that proton channels extrude the cytosolic acid generated by the consumption of NADPH during the respiratory burst; (2) VSOP/Hv1^−/− neutrophils were more depolarized than control neutrophils, as predicted from the lack of the compensating charge provided by proton channels. The cytosolic acidification is detrimental to the enzymatic activity of the phagocyte NADPH oxidase (Morgan et al., 2005), whereas the membrane depolarization hinders the flow of electrons across the flavocytochrome, which is voltage dependent (DeCoursey et al., 2003; Petheo and Demaurex, 2005). These two defects can account for the 80% reduction in H₂O₂ production that we observed in peripheral blood neutrophils stimulated with PMA. This inhibition is more pronounced than the 30% reduction reported by Okochi et al. (2009) in thioglycollate-activated peritoneal neutrophils and slightly higher than the 65% inhibition reported by Ramsey et al. (2009) in bone marrow neutrophils. The difference might be the result of the different assays used (cytochrome c vs. Amplex red) or might reflect differences between neutrophils from marrow, blood, and peritoneum. We also observed that Ca²⁺ influx was reduced by ∼60% in VSOP/Hv1^−/− neutrophils stimulated with PMA, most likely as a result of the membrane depolarization and cytosolic acidification. The reduced Ca²⁺ influx might hinder the oxidase in cells stimulated with chemoattractants, as in these conditions oxidase activity depends on the magnitude of the Ca²⁺ signal (Hallett et al., 1990).

The lack of proton currents in cells exhibiting normal electron currents also unambiguously establishes that the oxidase itself is not a proton channel. The notion that the oxidase contains a proton channel was postulated from thermodynamic considerations by Henderson et al. (1987), who provided several pieces of evidence for this mechanism. We did not support the hypothesis that the oxidase is a proton channel initially because cells from X-linked CGD patients lacking the gp91^phox molecule had normal proton currents (Nanda et al., 1994). We revived the Henderson hypothesis when we observed that cells with an active oxidase exhibit proton currents with remarkable gating properties (Bánfi et al., 1999). These novel currents were absent in cells from X-CGD patients, indicating that they required the gp91^phox molecule. The recognition that gp91^phox contains a proton channel motif (Starace et al., 1997) suggested that the flavocytochrome could function as a proton channel and led to the cloning of the NOX homologues (Bánfi et al., 2000). Heterologous expression of NOX homologues in HEK-293 and CHO cells generated proton currents that recapitulated the properties of endogenous proton currents (Bánfi et al., 2000; Maturana et al., 2001), validating the channel theory. However, proton currents were not observed in COS-phox cells expressing a functional oxidase (Morgan et al., 2002), suggesting instead that the oxidase modulates a proton channel. Our findings now provide conclusive evidence that gp91^phox is not a proton channel. Instead, VSOP/Hv1 is the only proton channel of phagocytes and its gating properties are enhanced when the oxidase is active (Musset et al., 2009). VSOP/Hv1 probably interacts with a component of the oxidase complex that assembles at the membrane upon phosphorylation, such as the p47^phox adaptor subunit. In agreement with this scenario, electron and proton currents cosegregate in excised patches (Petheö et al., 2003), suggesting that VSOP/Hv1 is closely associated with an oxidase component.

We also demonstrate that the lack of proton channels markedly reduces the influx component of calcium signals in neutrophils. This important secondary defect could be predicted from the increased depolarization that we observed in these cells when their oxidase was activated. A role for the oxidase in limiting calcium influx was previously reported (Demaurex et al., 1994; Geiszt et al., 1997; Rada et al., 2003; for review see Rada et al., 2005) but the ability of proton channels to sustain Ca²⁺ influx was not testable because the proton channel inhibitor Zn²⁺, like most divalent cations, also inhibits SOCE channels. In this paper, we show that ablation of the VSOP/Hv1 gene is associated with a ∼60% reduction in SOCE and with a ∼85% reduction in the calcium signal evoked by the chemoattractant fMIVIL. The Ca²⁺ defect was more apparent when the oxidase was preactivated with PMA, as predicted from the reduced driving force for calcium ions imparted by the depolarization, but was also observed in VSOP/Hv1^−/− neutrophils exposed to fMIVIL alone (Fig. 4 B). The increased cytosolic acidification might also hinder the activity of SOCE channels. This indicates that cells lacking VSOP/Hv1 proton channels cannot generate normal Ca²⁺ signals when their oxidase is pumping electrons across the plasma or phagosomal membrane. This signaling defect can have important consequences for the maturation of the phagosome and for the ability of neutrophils to kill microbes. Global Ca²⁺ elevations accelerate phagocytosis by activating calpain (Dewitt and Hallett, 2002), whereas local Ca²⁺ elevations around the phagosome are required for the fusion of lysosomes with phagosomes (Jaconi et al., 1990). The periphagosomal increase in Ca²⁺ is caused by the opening of SOCE channels on the membrane of phagosomes (Lundqvist-Gustafsson et al., 2000), small compartments which can depolarize significantly (Steinberg et al., 2007). Our observation that proton channel ablation depolarizes cells during oxidase activation suggests that phagosomes, where the oxidase normally assembles, are depolarized in cells lacking VSOP/Hv1. The loss of calcium influx at the phagosomal membrane is thus likely to impair cortical actin depolymerization. Accordingly, we observed a thicker ring of polymerized actin around zymosan particles ingested by VSOP/Hv1^−/− neutrophils. The thicker cortical actin cytoskeleton might prevent the docking of oxidase subunits to the phagosome and the fusion of lysosomes with phagosomes (Jaconi et al., 1990). Whether lack of VSOP/Hv1 proton channels impairs the fusion of granules to phagosomes remains to be established, but we document here that other Ca²⁺-dependent functions of neutrophils are altered. VSOP/Hv1^−/− neutrophils adhered more firmly to the substrate and migrated less efficiently when stimulated with the chemoattractant fMIVIL. Neutrophil migration was restored by the Ca²⁺ ionophore ionomycin, demonstrating that the calcium and migration defects are causally related. Thus, actin-based motility is altered in neutrophils lacking proton channels. The microbicidal activity of neutrophils in vivo requires their migration toward invading pathogens (chemotaxis), followed by the phagocytosis and killing of microbes, processes which are all orchestrated by calcium signals. In macrophages, inhibition of Ca²⁺ influx leads to loss of leading-edge PI3K activity, disassembly of F-actin, cessation of ruffling, and decay of chemoattractant signals (Evans and Falke, 2007). Defective bacterial killing was recently reported in VSOP/Hv1^−/− neutrophils and attributed to decreased superoxide production (Ramsey et al., 2009). Our findings not only explain the decreased superoxide production, but indicate that the failure of VSOP/Hv1^−/− neutrophils to kill bacteria might also be a result of the altered pH homeostasis and Ca²⁺ signaling of these cells, which impair their mobility and phagocytic ability. The loss of VSOP/Hv1 proton channels thus has multiple detrimental consequences on the bactericidal activity of neutrophils.

In summary, we show that proton channels sustain superoxide production and calcium entry in neutrophils by preventing membrane depolarization and cellular acidification. VSOP/Hv1 is the only proton channel of neutrophils. Its deletion aborts calcium signals and inhibits neutrophil motility and, probably, also the docking of granules and oxidase subunits with phagosomes, thereby impairing the killing of microbes by neutrophils.

fMIVIL was synthesized at UNIGE Peptides Synthesis Platform. This pentapeptide from Listeria monocytogenes was used instead of N-formyl-Met-Leu-Phe (fMLF) because it is ∼100-fold more potent than fMLF in activating mouse neutrophils (Southgate et al., 2008). Other chemicals were purchased from Sigma-Aldrich unless indicated otherwise.

Mice bearing a targeted disruption in the VSOP/Hv1 (VSOP/Hv1^−/−, backcrossed eight times) were previously described (Okochi et al., 2009). WT mice (VSOP/Hv1^+/+) were of the same genetic background (C57BL6J). Animals enrolled for the study were aged between 1 and 6 mo. All animal husbandry and experiments were approved by and performed in accordance with guidelines from the animal research committee of the University of Geneva.

Mouse blood (350 ± 50 µl per animal) was collected from 13 VSOP/Hv1^+/+and 14 VSOP/Hv1^−/− mice in EDTA-tubes by tail bleeding. Blood was collected every 2 wk from the same animals, which were bled four times on average (total 54 bleedings). The exact number of mice used for each experiment is given in the figure legends. NaCl (0.9%) was added to a final volume of 4 ml. After centrifugation at 2,000 rpm for 10 min, cells were resuspended in 1 ml HBSS (without calcium, magnesium, and phenol red) supplemented with 0.2% EDTA. The cells were overlaid afterward on a three-layer Percoll density gradient 72, 63, and 50%, respectively, diluted in HBSS (100% Percoll = nine parts Percoll and one part 10× HBSS) and centrifuged at 3,000 rpm for 25 min without breaking. Neutrophils were harvested from the 63/72% interface after carefully removing the cells from the upper phases. After one wash, remaining red cells in the neutrophils fraction were eliminated by hypotonic lysis (0.5 ml water for 35 s and 0.5 ml 1.8% NaCl). After a final wash, 100,000–500,000 cells were obtained per mouse. Neutrophils were kept in Tyrode solution containing 140 mM NaCl, 5.4 mM KCl, 1.8 mM MgCl₂, 1.8 mM CaCl₂, 10 mM Hepes, and 10 mM glucose, pH 7.2, and used within 6 h.

Mice were sacrificed and femur and tibia from both hind legs were removed and cleared of all remaining soft tissue attachments. Then, the extreme distal tip of each extremity was cut off and the bones were flushed with HBSS medium (with calcium and magnesium) containing 0.5% BSA, 1% glucose (Sigma-Aldrich), and 20 mM Hepes, pH of 7.2, using a 25G 5/8” needle. After bone marrow was flushed on ice, marrow was centrifuged at 1,300 rpm for 10 min and a hypotonic lysis was directly performed afterward to eliminate red blood cells. The remaining cells were concentrated in 3 ml HBSS medium and treated on a three-layer Percoll density gradient exactly as described in the previous section. The total cell number obtained with this protocol was 5–8 × 10⁶ cells. In total, 11 VSOP/Hv1^−/− and 11 VSOP/Hv1^+/+ mice were sacrificed. Blood and bone marrow cells preparations contained >90% neutrophils as determined by visual counting. Neutrophils viability was >95% as assessed by Trypan blue exclusion.

Both the whole-cell and perforated patch configuration of the patch-clamp technique were used to record proton and electron currents via a patch-clamp amplifier (Axoclamp 200B; MDS Analytical Technologies) interfaced to a personal computer. Pipettes were made from GC150F-10 Harvard Apparatus borosilicate glass capillaries using a PC-10 Narishige puller. Patch electrodes of 3–8 MΩ were filled, for whole-cell recordings, with an internal solution containing: 118 mM/liter NMDG (N-methyl-d-glucamine), 63 mM/liter TMAMeSO₃ (tetramethylammonium methanesulfonate), 1 mM/liter MgCl₂, 119 mM/liter MES, and 1 mM/liter EGTA. This solution was adjusted to pH_i 6.1 with TMAOH. Extracellular solution contained 140 mM/liter TMAMeSO₃, 1.8 mM/liter CaCl₂, 100 mM/liter Hepes, and 40 mM mannitol and adjusted at pH_o 7.2. The osmolarities of both external and internal solutions were maintained at 300 and 290 mOsm, respectively. For perforated patch recordings, the pipette solution contained 130 mM/liter TMAMeSO₃, 25 mM/liter (NH₄)₂SO₄, 2 mM/liter MgCl₂, 10 mM/liter Hepes, 1 mM/liter EGTA, and 0.5 mg/ml nystatin. Extracellular solution contained 130 mM/liter TMAMeSO₃, 25 mM/liter (NH₄)₂SO₄, 2 mM/liter MgCl₂, 1.5 mM/liter CaCl₂, 10 mM/liter Hepes, 1 mM/liter EGTA and 10 mM/liter glucose. The pH of both solutions was adjusted at 7.0. No liquid junction potential correction was applied and all experiments were performed at room temperature (20–25°C).

Neutrophils were loaded with 2 µM BCECF/AM for 20 min, seeded on 25-mm glass coverslips that were inserted in a thermostatic chamber (Harvard Apparatus), and imaged on an Axiovert S100 TV through a 40× 1.3 NA oil-immersion objective (Carl Zeiss, Inc.) using a cooled 16-bit charge-coupled device back-illuminated frame transfer camera (MicroMax; Roper Industries). Cells were alternately excited at 490 and 440 nm with a monochromator (DeltaRam; Photon Technology International) and images acquired using a 535 ± 35-nm emission filter (Omega Optical). Analysis was performed with the MetaFluor 6.2 software (MDS ANalytical Technologies). To exclude any contribution of the Na⁺/H⁺ antiporter, experiments were performed in Na⁺-free medium containing: 140 mM/liter NMDG-Cl, 5 mM/liter KCl, 1 mM/liter MgCl₂, 1.8 mM/liter CaCl₂, 10 mM/liter Hepes, and 10 mM/liter glucose, pH 7.4. For calibration, 5 µg/ml nigericin and 5 µM monensin (Sigma-Aldrich) were added to solutions containing 140 mM/liter KCl, 1 mM/liter MgCl₂, 0.2 mM/liter EGTA, and 20 mM/liter Hepes, pH 7.0–7.5, or MES, pH 6.0–6.5.

Neutrophils were assayed in HBSS medium with a cell density of 20,000 cells/well in a 96-well flat-bottom plate in a 200-µl final volume. For H₂O₂ measurements, cells were incubated in the presence of 25 µM Amplex red and 0.005 U/ml horseradish peroxidase. The inhibitors Zinc (1 mM) and DPI (10 µM) were added to the wells 10 min before the recordings. 0.1 µM PMA was added at time t = 0 and fluorescence measured at 590 nm on a FLUOstar (BMG Labtech) microplate reader every minute for 1 h at 37°C. Each condition was performed in triplicate. Six different concentrations of H₂O₂ were used to obtain a calibration curve. The fMIVIL effect on ROS production was measured with the chemiluminescent probe 8-amino-5-chloro-7-phenylpyrido[3,4-d]pyridazine-1,4(2H,3H)dione (L-012) (Imada et al., 1999). Chemiluminescence was measured every 8 s during 15 min on the FLUOstar microplate reader.

The voltage-sensitive bis-oxonol fluorescent dye DiBAC(4)3 was used to study membrane potential changes of neutrophils. Cells were seeded on 25-mm glass coverslips and the acquisition started directly after loading the cells with 250 nM of the dye in Tyrode medium. Cells were excited at 500 nm through a 430/500/575-nm triple band beamsplitter (Chroma Technology Corp.) and images acquired at 535 ± 30 nm.

Neutrophils were loaded with 2 µM Fura-2/AM for 20 min. When indicated, cells were activated by adding 0.1 µM PMA 10 min before starting the acquisition. Fura-2 fluorescence was imaged using alternate excitation at 340 and 380 nm and 510 ± 40 nm emission.

Bone marrow neutrophils were seeded in a Greiner 96-well flat-bottom plate at 500,000 cells/well in HBSS medium supplemented with 0.5% BSA, 1% glucose, and 20 mM Hepes. Neutrophils were prewarmed at 37°C. The wells were coated with 0.5% BSA and the plate was preheated for 15 min at 37°C. Agonists were added to each well (200 µl/well) and the adhesion and chemokinesis of neutrophils imaged with an ImageXpress Micro (MDS Analytical Technologies) automated fluorescence imaging system. Data were analyzed with the MetaXpress software (MDS Analytical Technologies). Images were taken every 15 s during 45 min.

Fig. S1 shows actin polymerization in VSOP/Hv1^−/− neutrophils. Fig. S2 shows the effect of fMIVIL on ROS production. Video 1 shows VSOP/Hv1^+/+ neutrophil migration upon fMIVIL stimulation. Video 2 shows the slower migration of neutrophils lacking VSOP/Hv1.

The authors would like to thank C. Castelbou for his excellent technical assistance.

This research was supported by grant 3100A0-118393 from the Swiss National Science Foundation (to N. Demaurex) and by grants from the Japan Ministry of Education, Culture, Sports, Science and Technology (to Y. Okamura).

The authors have no conflicting financial interests.