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Journal of Physiology (2001), 535.3, pp. 767-781
© Copyright 2001 The Physiological Society
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ABSTRACT |
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INTRODUCTION |
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The NADPH oxidase enzyme complex is a critically important component of the host defence system against bacterial invaders. In resting phagocytes NADPH oxidase is inactive, with its several protein components partitioned between the cytosolic and membrane compartments (Babior, 1999). The active enzyme is assembled in the membranes of phagocytes upon cell activation by phagocytic stimuli such as opsonized bacteria or by soluble stimuli such as chemotactic peptides and the artificial stimulus phorbol myristate acetate (PMA). The active NADPH oxidase complex generates the superoxide anion (O2-) and other reactive oxygen intermediates that are toxic to micro-organisms. The importance of these reactions is evident in chronic granulomatous disease, in which the absence of a functioning NADPH oxidase is manifested as a lethal susceptibility to recurrent bacterial infection if left untreated (Babior, 1999).
NADPH oxidase is electrogenic, transporting electrons out of the cell in order to reduce O2 to O2-, and compensatory electrogenic proton efflux is activated during the respiratory burst (Henderson et al. 1987, 1988a,b; Nanda & Grinstein, 1991). Henderson et al. (1987, 1988a,b) postulated that this H+ efflux was mediated by voltage-gated proton channels. In support of this hypothesis, voltage-gated proton currents have been found in every phagocyte or related cell line that has been studied (Eder & DeCoursey, 2001), but many questions remain. Although NADPH oxidase-associated electron flux and H+ efflux are both activated by PMA, a potent agonist for the respiratory burst, it is not clear whether the H+ conductance (gH) is activated independently by PMA or indirectly as a consequence of the oxidase functioning. Both the decrease in pHi and the depolarization of the membrane potential that tend to occur when the oxidase functions can activate voltage-gated proton channels. A role for diffusible intracellular second messengers in the activation of both electron and proton currents in human neutrophils is suggested by the observation that PMA is ineffective in whole-cell studies but activates both electron and proton currents in the permeabilized-patch configuration (DeCoursey et al. 2000). The oxidase inhibitor diphenylene iodinium (DPI) inhibited the electron currents, which reflect NADPH oxidase activity, but not the increased H+ current amplitude in neutrophils, indicating dissociation between the operation of these two conductances. However, the PMA-induced slowing of the closing rate of H+ channels was reversed by DPI, suggesting that some functional link exists between NADPH oxidase and voltage-gated proton channels (DeCoursey et al. 2000).
Like neutrophils, eosinophils play an important role in host defence and are also implicated in the pathogenesis of a number of disorders, including most notably asthma and other allergic inflammatory disorders (Rothenberg, 1998; Gleich, 2000). The composition of the NADPH oxidase system in eosinophils is similar to that in neutrophils and other phagocytes (Someya et al. 1997; Giembycz & Lindsay, 1999), and generation by this enzyme of O2- and other oxygen radicals contributes to both the host defence and pathogenic activities of eosinophils (Wu et al. 2000). Indeed, eosinophils exhibit a particularly vigorous respiratory burst compared with neutrophils and other phagocytes (DeChatelet et al. 1977; Yamashita et al. 1985; Shult et al. 1985; Petreccia et al. 1987; Yagisawa et al. 1996; Someya et al. 1997), with consequently larger electron currents (Schrenzel et al. 1998; DeCoursey et al. 2000). Their vigorous respiratory burst and the existence of techniques that allow their ready isolation (Hansel et al. 1991) make the human eosinophil an attractive cell for the study of NADPH oxidase function.
It was proposed recently that eosinophils have two distinct varieties of voltage-gated proton channels, one that is present in resting cells, and another with different properties that is activated during the respiratory burst (Bánfi et al. 1999). We named these channels type p and type x respectively, for 'phagocyte' and 'oxidase' (Eder & DeCoursey, 2001). However, several types of evidence suggest that there is a single type of voltage-gated proton channel in neutrophils, whose properties are modulated strongly when NADPH oxidase is activated by PMA (DeCoursey et al. 2000). The voltage-gated proton conductance that is activated during the respiratory burst in human eosinophils resembles that activated in human neutrophils (DeCoursey et al. 2000), and the data are consistent with the idea that a single population of H+ channels exists in human eosinophils or neutrophils.
A preliminary account of this work has been published (Cherny et al. 2001a).
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METHODS |
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Eosinophil and neutrophil isolation
Venous blood was drawn from healthy adult volunteers under informed consent according to procedures approved by our Institutional Review Board and in accordance with Federal regulations. Neutrophils were isolated by density gradient centrifugation as described previously (Haskell et al. 1995) with one modification. In the two cycles of hypotonic lysis performed for removal of the red blood cells, isotonicity was restored by the addition of 2 
concentrated Hanks' balanced salt solution (HBSS; cat. no. 14185, Gibco, Grand Island, NY, USA) (without Ca2+ or Mg2+) containing 5 mM Hepes, and adjusted to pH 7.4. An aliquot of the neutrophil preparation was held on ice, and eosinophils were isolated from the neutrophil preparation by negative selection using anti-CD16 immunomagnetic beads (Hansel et al. 1991) as described by the manufacturer (Miltenyi Biotec Inc., Auburn, CA, USA). The eosinophils or neutrophils were suspended in Hepes (10 mM)-buffered HBSS (with Ca2+ and Mg2+, Gibco cat. no. 14065), adjusted to pH 7.4, containing 1 mg ml-1 human serum albumin (Hepes-HBSS-HSA buffer). Eosinophil purity was routinely greater than 98 % as determined by counting Wright-stained cytospin preparations. Neutrophil purity was routinely 95 % or greater. Cells from at least 14 different donors were used in this study.
Superoxide anion production
Superoxide anion (O2-) production was measured essentially as described elsewhere (Horie & Kita, 1994). Briefly, eosinophils or neutrophils (each at 2.5 105 cells ml-1) were incubated with 8 nM PMA in Hepes-HBSS-HSA buffer containing 50 µM cytochrome c for 45 min at 37 °C. Incubations were performed in a flat-bottomed 96-well tissue culture plate (Costar, Acton, MA, USA) in a Ceres UV900HDi microplate reader (Bio-Tek Instruments, Inc., Winooski, VT, USA), and absorbance at 550 nm was recorded at 5 min intervals. Tissue culture wells were precoated with human serum albumin (Horie & Kita, 1994). Total incubation volume was 0.2 ml. Production of O2- was calculated as described previously (Horie & Kita, 1994). Results are expressed in nanomoles O2- per 105 cells after subtraction of spontaneous production, which was measured in the absence of PMA stimulus.
Electrophysiology
We studied freshly isolated eosinophils and eosinophils incubated overnight at 37 °C in RPMI 1640 medium containing 25 mM Hepes and L-glutamine (Gibco), supplemented with 10 % fetal bovine serum (Bio-Whittaker, Walkersville, MD, USA), 100 units ml-1 penicillin, 100 µg ml-1 streptomycin (Sigma), and 1 ng ml-1 recombinant human GM-CSF (R & D Systems, Inc., Minneapolis, MN, USA). No difference between the properties or responses of freshly isolated and incubated eosinophils was detected.
For permeabilized-patch recording, the pipette and bath solutions usually contained 50 mM NH4+ in the form of 25 mM (NH4)2SO4, 2 mM MgCl2, 5 mM Bes or Pipes buffer, 1 mM EGTA, and were adjusted to pH 7.0 with TMAOH. We added ~500 µg ml-1 solubilized amphotericin B (~45 % purity) (Sigma) to the pipette solution, after first dipping the tip in amphotericin-free solution.
Micropipettes were pulled using a Flaming-Brown automatic pipette puller (Sutter Instruments, San Rafael, CA, USA) from 7052 glass (Garner Glass Co., Claremont, CA, USA), coated with Sylgard 184 (Dow Corning Corp., Midland, MI, USA), and heat polished to a tip resistance ranging typically between 3 and 10 M
. Electrical contact with the pipette solution was achieved by a thin sintered Ag-AgCl pellet (In Vivo Metric Systems, Healdsburg, CA, USA) attached to a Teflon-encased silver wire, or simply a chlorided silver wire. A reference electrode made from a Ag-AgCl pellet was connected to the bath through an agar bridge made with Ringer solution (containing (mM): 160 NaCl, 4.5 KCl, 2 CaCl2, 1 MgCl2, 5 Hepes, adjusted to pH 7.4). The current signal from the patch clamp (EPC-7 from List Electronic, Darmstadt, Germany, or Axopatch 200B from Axon Instruments, Foster City, CA, USA) was recorded and analysed using an Indec Laboratory data acquisition and display system (Indec Corporation, Sunnyvale, CA, USA) or pCLAMP software supplemented by Microsoft Excel and SigmaPlot (SPSS Inc., Chicago, IL, USA). Seals were formed with Ringer solution in the bath, and the potential zeroed after the pipette was in contact with the cell. For the experimental set-up where most measurements were done, the bath temperature was kept at 20-21 °C by Peltier devices and monitored by a resistance temperature detector (RTD) element (Omega Scientific, Stamford, CT, USA) in the bath. For the other experimental set-ups, data were collected at room temperature (20-25 °C).
Heterogeneous responses to stimulation and possible effects of adherence
There was variability in the behaviour of unstimulated eosinophils and in their response to PMA. Within the first ~10 min of establishing a seal, the proton current (IH) tended to increase and often the reversal potential (Vrev) shifted from a positive value to near 0 mV. IH would be expected to increase over time as the amphotericin in the pipette inserts into the membrane and establishes control over the membrane potential. Most measurements were done with a symmetrical NH4+ gradient, nominally setting pHi to 7.0, which is lower than is likely to occur in an untreated eosinophil. IH would tend to increase as pHi decreases. We started to make measurements after the H+ currents exhibited stable behaviour.
In the absence of any added stimulus some cells exhibited signs of 'activation': slow tail current decay, activation at voltages near or negative to the proton equilibrium potential (EH), and in some cases inward current (presumed electron current) at a holding potential (Vh) of -60 mV. These signs were present immediately, developed over time, or disappeared over time. Cells displaying spontaneous activation responded to DPI with speeding of the tail current time constant (
tail) and block of inward electron current at -60 mV (Cherny et al. 2001b, Fig. 2), confirming that the spontaneously active state included both NADPH oxidase activity and the attendant changes in H+ current properties associated with PMA activation. As was found in neutrophils (DeCoursey et al. 2000), DPI had no effect on non-activated eosinophils, H+ currents or the leak current.
In early studies, we noticed that eosinophils that were allowed to settle onto the glass chamber rapidly became attached and began to spread. Our impression was that these adherent cells were more likely than non-adherent cells to appear 'activated' (according to the criteria listed in the previous paragraph) as soon as recording had begun. Because they seemed less predisposed to adhere to albumin-coated glass, eosinophils were placed above coverslip fragments precoated by incubation with 20 % bovine serum albumin (Sigma). Although this precaution appeared to reduce spontaneous activation, some cells still displayed signs of activation. In many cells, these signs disappeared within a few minutes, after which the cell was capable of responding to a challenge with PMA.
Bath solution changes
Solutions and physiologically active substances were applied by flushing the entire bath solution (~400 µl volume) with ~10 bath volumes, which took ~10-20 s. The time of a bath change was taken as roughly the time the first volume replacement occurred. We did not perfuse the chamber continuously. As a control for mechanically induced responses, the bath solution was usually first replaced with an identical solution before adding a test solution. Changing the bath solution transiently increases H+ currents due to a small increase in temperature. Due to the strong temperature sensitivity of the H+ current (Kuno et al. 1997; DeCoursey & Cherny, 1998), even an ~1 °C change has a definite effect.
Conventions
Currents and voltages are presented in the normal sense: upward currents represent positive charge flowing outward through the membrane, and potentials are expressed relative to the potential of the original bath solution which is defined as 0 mV. Current records are presented without correction for leak current or liquid junction potentials.
Data analysis
The time constant of H+ current activation (act) was obtained by fitting the current record by eye with a single exponential after a brief delay (DeCoursey & Cherny, 1995):
I(t) = (I0 - I
)exp(-(t - tdelay)/act), (1)
where I0 is the initial amplitude of the current after the voltage step, I is the steady-state current amplitude, t is the time after the voltage step, and tdelay is the delay. The H+ current amplitude is (I0 - I). No other time-dependent conductances were observed consistently under the ionic conditions employed. Tail current time constants (tail) were obtained by fitting tail currents to a single exponential:
I(t) = I0exp((-t/tail) + I), (2)
where I0 is the amplitude of the decaying part of the tail current.
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RESULTS |
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Properties of voltage-gated proton currents in unstimulated eosinophils in the permeabilized patch configuration
Eosinophils were studied using the permeabilized-patch approach in order to preserve intracellular second messenger pathways. We established an NH4+ gradient in order to 'clamp' pHi (Grinstein et al. 1994; DeCoursey et al. 2000). In most experiments the NH4+ gradient was symmetrical (50 mM NH4+ in bath and pipette solutions), which kept the pHi near pHo (7.0 unless otherwise stated). Under these conditions, most human eosinophils exhibited robust voltage-gated proton currents, as illustrated in Fig. 1A. The average H+ current measured at the end of an 8 s pulse to +80 mV was 48 ± 31 pA (mean ± S.D.) in a sample of 16 consecutively studied eosinophils. The H+ selectivity of these currents was confirmed by measuring the tail current reversal potential (Vrev) as illustrated in Fig. 1B. In this experiment, the tail currents reversed near 0 mV, consistent with the symmetrical NH4+ gradient. When pHi was varied indirectly by changing the NH4+ concentration in the bath solution Vrev changed in reasonable agreement with the Nernst potential for NH4+, which ideally establishes an identical H+ gradient. In one to four measurements at each [NH4+]o in three cells studied with 50 mM NH4+ in the pipette solution, Vrev averaged -32 mV at 15 mM NH4+ (EH = -30 mV), -44 mV at 9 mM NH4+ (EH = -43 mV), -70 mV at 3 mM NH4+ (EH = -71 mV), and -82 mV at 1 mM NH4+ (EH = -99 mV). Thus, the time-dependent currents are H+ selective and the applied NH4+ gradient is adequate to establish pHi, as was previously demonstrated using this approach in human neutrophils (DeCoursey et al. 2000).
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Figure 1. Properties of voltage-gated proton currents in unstimulated human eosinophils studied using the permeabilized patch technique A, a family of currents during 8 s voltage pulses (at 30 s intervals) from -40 mV through +80 mV in 20 mV increments from Vh = -60 mV. B, tail currents measured in the same cell. A 4 s prepulse to +80 mV opened H+ channels, then the potential was stepped to -30 mV through +20 mV in 10 mV increments. Vrev is close to 0 mV, as expected for a symmetrical pH gradient (50 mM NH4+ in pipette and bath solutions, pH 7.0). Low-pass filtered at 100 Hz. C, mean ± S.D. time constants of activation, | ||
Gating kinetics
The kinetics of H+ current activation during a depolarizing pulse was quantified by fitting a single exponential after a delay. During large depolarizations the H+ current in some cells continued to rise progressively at a slower rate. These currents could be fitted arbitrarily with an exponential plus a linearly rising component. We adopted the single exponential approach rather than introducing a more complex function for three reasons: (1) it is simple; (2) we have used it previously in a number of cell types; and (3) as a result of the high H+ current density in eosinophils, diffusion artifacts may alter the apparent kinetics.
The activation time constant (act) obtained in this way is plotted in Fig. 1C (
). Activation of H+ current was faster at more positive potentials, but with modest voltage dependence, with act changing e-fold in 59 mV. The time constant of deactivation of the H+ current (tail) was obtained by fitting tail currents like those illustrated in Fig. 1B with a single exponential. A single exponential provided a good fit over the entire voltage range negative to the threshold for activating the H+ conductance (gH). The voltage dependence of deactivation was almost twice as steep as that of activation, with tail changing e-fold in 32 mV (Fig. 1C, 
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Effects of PMA on H+ currents
Although PMA has no effect on H+ currents in phagocytes studied in whole-cell configuration, profound effects are observed in permeabilized-patch recording conditions (DeCoursey et al. 2000). Figure 2A and B illustrates families of voltage-gated proton currents recorded in the same eosinophil before and after addition of PMA. After PMA stimulation, the H+ current (IH) is larger and the gH was activated at substantially more negative voltages. The NADPH oxidase inhibitor DPI had little immediate effect on H+ currents in PMA-stimulated cells (Fig. 2C), except that it accelerated tail current decay. IH was somewhat larger after DPI treatment in this cell, but this effect was not always seen.
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Figure 2. Effects of PMA and DPI on H+ current properties A, currents recorded in an untreated eosinophil during depolarizing pulses from Vh = -60 mV to -40 mV through +80 mV in 20 mV increments are superimposed. B, currents in the same cell after addition of 30 nM PMA. Pulses were applied in 10 mV increments. C, currents in the same cell after addition of 3 µM DPI (and washout of PMA). The arrows indicate the level of zero current. D, chord conductance-voltage relationships in the same eosinophil as in A-C, before ( | ||
Activation of voltage-gated proton channels in unstimulated eosinophils occurred at potentials more positive than +20 mV (Fig. 2D). After PMA treatment, the gH-V relationship shifted toward substantially more negative voltages. We quantified this shift by estimating the threshold voltage at which time-dependent currents were first elicited (Vthreshold), or after which distinct tail currents were seen upon repolarization to -60 mV. The average shift in Vthreshold produced by PMA in eosinophils by these criteria was -42.6 ± 1.8 mV (mean ± S.E.M., n = 23). The addition of DPI did not change Vthreshold much, although a slight depolarizing shift was sometimes evident. Often IH increased somewhat, as in Fig. 2D, but it is not clear whether this was a genuine effect of DPI or simply the continuation of progressive enhancement by PMA. Long after PMA exposure in some cells there was a return toward control values of all parameters. In these cells, the gH-V relationship gradually shifted toward more positive voltages, IH became smaller, act became slower, and tail became faster.
Because the gH-V relationship shifted toward substantially more negative voltages after PMA treatment, the threshold often occurred at potentials more negative than Vrev, which resulted in the activation of inward H+ currents. Inward H+ currents are not normally observed in cells studied in whole-cell recording conditions (Eder & DeCoursey, 2001), but occur in eosinophils under conditions that activate NADPH oxidase (Bánfi et al. 1999). These currents are small because the driving force near Vrev is small; however, they can be seen at high gain (data not shown), and can be confirmed by examination of the tail currents upon repolarization to -60 mV.
The effect of PMA on act is shown in Fig. 2E. The average values of act measured at several voltages before () and after PMA (
) are plotted. To a first approximation, the activation rate increased uniformly at all voltages. Also plotted is the average act in cells that eventually reverted to control-like behaviour (
).
Shifts of Vrev
Because NADPH oxidase generates intracellular protons, its activation will tend to lower pHi. In our experiments, this tendency is opposed by the applied NH4+ gradient, but the additional acid load on this system may still result in a lower pHi. Indeed, Vrev measured after PMA treatment tended to be more negative than in the same cells before PMA. The average change in Vrev in 16 cells measured before and after exposure to 30-60 nM PMA was -5.8 ± 1.5 mV (mean ± S.E.M.), indicating that slight cytoplasmic acidification (~0.1 pH unit) did occur in spite of the NH4+ gradient. Cytoplasmic acidification due to the activity of NADPH oxidase should be reversed by DPI. In seven cells in which Vrev was measured after PMA and again after DPI, the average change in Vrev was +3.0 ± 0.8 mV.
Superoxide anion release by eosinophils induced by PMA
Voltage-gated proton channels are believed to play a role during the respiratory burst in eosinophils and other phagocytes (Henderson & Chappell, 1996; DeCoursey & Grinstein, 1999). We therefore examined superoxide production in response to PMA stimulation in eosinophils. Figure 3A shows the time course of PMA-stimulated superoxide production by neutrophils () and eosinophils (
) from four donors. Consistent with previous studies (DeChatelet et al. 1977; Yamashita et al. 1985; Shult et al. 1985; Petreccia et al. 1987; Freiburghaus et al. 1991; Yagisawa et al. 1996; Someya et al. 1997), the respiratory burst was larger and faster in eosinophils than in neutrophils. The difference in the time course is more apparent when the rate of O2- release is plotted, as in Fig. 3B. In both cell types, the peak O2- production occurred 5-10 min after PMA addition. In eosinophils, the respiratory burst was transient, with no O2- released after 25 min. In neutrophils release persisted longer and was still ~40 % the maximal rate 40 min after PMA stimulation. The peak rate in eosinophils was 3.2 times greater than in neutrophils.
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Figure 3. Time course of PMA-stimulated O2- release from human eosinophils and human neutrophils A, comparison of net cumulative O2- release from human eosinophils ( | ||
Time course of PMA effects
Figure 4A illustrates the time course of PMA effects on proton and electron currents in an eosinophil. Pulses were applied from Vh = -60 mV to a test potential of +60 mV at 30 s intervals. Characteristically, IH increased after a delay. In Fig. 4A, no effect was observed 30 s after addition of 30 nM PMA, and the maximal rate of change occurred 1-2 min after stimulation. In addition to increasing IH substantially, PMA stimulation caused the H+ current activation time course to become faster (smaller act). Finally, the rate of tail current decay (the rate that H+ channels close upon repolarization to -60 mV) became slower (tail increased).
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Figure 4. Time dependence of the electrophysiological effects of PMA on human eosinophils in the permeabilized-patch configuration A, current records during test pulses to +60 mV from a holding potential of -60 mV are superimposed. The dashed line shows the zero current level. Two control currents (labeled '0') were recorded before and after a bath change to control for possible flow artifacts. After application of 30 nM PMA test pulses were applied every 30 s; the first 7 are shown here. The first current recorded after PMA addition is superimposed on the control currents. IH increased noticeably 1 min after PMA addition and increased progressively over the next several minutes. The largest current illustrates a quasi-steady-state reached ~6 min after PMA addition. The inward current at -60 mV (electron current) increased progressively after PMA addition, with the first and last record labelled according to the time after PMA addition (0 and 6 min). The tail currents increased in amplitude and became progressively slower. Low-pass filter 100 Hz. B, the changes in various parameters during exposure of another eosinophil to PMA and then DPI are plotted. The abscissae indicate roughly the time elapsed after establishing permeabilized patch recording. The dashed vertical lines indicate the time the bath solution was changed to introduce PMA or DPI, as indicated. H+ current parameters are IH ( | ||
In addition to its effects on the H+ conductance, PMA usually activated an inward current in eosinophils. The progressive increase in the inward current at -60 mV after PMA stimulation is evident in Fig. 4A and also can be seen in Fig. 2 (note arrows indicating zero current). As shown in Fig. 4B (+), this current was inhibited by DPI, an inhibitor of NADPH oxidase (Robertson et al. 1990). The inward current therefore reflects the efflux of electrons across the membrane mediated by NADPH oxidase and is termed electron current (Ie). The Ie amplitude was on average -6 pA (Table 1), which is several times larger than in neutrophils studied under similar conditions (DeCoursey et al. 2000). In contrast to the H+ current, which can be observed only by applying a depolarizing voltage pulse to open voltage-gated proton channels, Ie is not voltage gated. Thus, it can be observed at -60 mV where there is no H+ current. This lack of voltage- and time-dependent gating of Ie allows clear separation from IH and enables both to be monitored in the same cell at essentially the same time.
Figure 4B shows the time course of the changes in H+ currents produced by PMA. Shortly after addition of PMA, activation (
) of H+ currents became faster, the IH amplitude () increased, and deactivation () of H+ currents became slower. Distinct inward electron current (Ie, +) appeared after a delay of 1-2 min. Addition of DPI turned off most of the PMA-induced inward current, supporting its origin as electron flux through the NADPH oxidase complex. DPI also largely reversed the slowing of tail produced by PMA, but DPI did not reverse the enhanced IH or the faster act. These effects of PMA are very similar to those reported previously in human neutrophils (DeCoursey et al. 2000). It should be emphasized, however, that there was considerable heterogeneity in the kinetics and magnitude of eosinophil responses to PMA. A remarkable result seen in this and most other experiments is that the time courses of changes in tail and Ie paralleled each other.
The time courses of the changes in the electrophysiological parameters are summarized in Table 1. Although the effects are quite similar to those described recently in human neutrophils studied under nearly identical conditions (DeCoursey et al. 2000), the response of eosinophils was slightly faster. As in neutrophils, the most rapid response was an increase in the rate of H+ current activation. In Fig. 4B, act was near its steady-state value before much Ie had turned on.
Spontaneous reversal of PMA effects
The PMA responses just described occurred in most eosinophils (42/51 or 82 %). The changes in all four parameters, however, reversed spontaneously, at least to some extent, in about half the cells that survived at least 20 min after PMA treatment. Figure 5 illustrates one such cell. PMA had its usual effects on H+ and electron currents, but before DPI was added, Ie had almost entirely disappeared. In parallel, the profoundly slowed tail returned to near its original value. It is noteworthy that in this cell in which tail had spontaneously recovered almost completely, addition of DPI had little effect and did not decrease tail beyond its original value. In contrast, in Fig. 4B DPI accelerated tail. Evidently, DPI reverses the slowing of tail induced by PMA, but this effect is not mediated by a direct pharmacological action on H+ channels. In other cells in which Ie turned off spontaneously, the PMA-induced slowing of tail usually reversed in parallel. In the experiment in Fig. 5, the IH amplitude and act returned to near their original values after DPI was added, but it is not clear whether the addition of DPI was responsible for this reversal. In other experiments, such as in Fig. 4B, DPI did not reverse these two parameters. In those cells in which the enhanced IH and the faster act returned toward control values, this recovery seemed to occur independently of the presence or absence of DPI.
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Figure 5. An eosinophil exhibiting a transient response to PMA Time course of electrophysiological changes produced by PMA in an eosinophil, with the same symbol definitions as in Fig. 4B. Ie (+) peaked ~3 min after PMA addition and then spontaneously turned off almost completely. Intriguingly, the slowing of | ||
As in the example in Fig. 5, the electron currents stimulated by PMA in many eosinophils were noticeably more transient than in neutrophils. Because electron current is not distinguishable from non-specific leak current unless it is shown to be inhibited by DPI, the entire time course of the response cannot be determined unambiguously. However, in a number of eosinophils, presumed Ie appeared within a few minutes of PMA stimulation, and then disappeared spontaneously within 5-10 min. We believe that the inward current activated by PMA in the eosinophil shown in Fig. 5 was Ie, because it occurred in synchrony with the changes in H+ currents that occurred during the PMA response. By contrast, the Ie in neutrophils declined spontaneously over time only infrequently (DeCoursey et al. 2000). We cannot absolutely rule out the possibility that in some cells H+ current parameters 'recovered' due to spontaneous patch rupture resulting in whole-cell configuration. However, the gradual time course of recovery and the survival of the cell in Fig. 5 with no increase in leak current suggests that spontaneous transition to whole-cell configuration in this cell is unlikely.
In Fig. 5, PMA was added for a second time after the electrophysiological parameters had returned to near their initial (unstimulated) values. There was no response. In other cells, repeated application of PMA sometimes produced a response, but the response was usually weaker than the initial response. During some secondary PMA exposures Ie was activated and IH increased, but the changes in gating kinetics of H+ currents were seen less often. We suspect that the durations of PMA and DPI exposures as well as the extent to which various parameters had returned toward normal influenced whether a secondary response occurred. It is noteworthy that most cells that appeared by various criteria to be activated spontaneously early in the experiment (see Methods, Heterogeneous responses to stimulation and possible effects of adherence) responded to PMA after returning to 'normal' behaviour. The multiplicity of events that occur during the first few minutes of an experiment (prodding the cell with the glass pipette tip, applying the NH4+ gradient to set pHi, and establishing voltage-clamp control during insertion of amphotericin into the membrane) obscures the precise state of the cell at that time. It is conceivable that every cell is 'activated' to some extent, and that most recover spontaneously before we have made the first measurement. In summary, secondary exposure of an eosinophil to PMA produced either no response or a weaker response, suggesting that some step in the PMA response pathway is not completely reversible within the time frame of these experiments.
Zinc sensitivity
All voltage-gated proton channels are inhibited by ZnCl2 (DeCoursey & Cherny, 1994). Zn2+ does not simply block H+ channels; it alters gating, slowing activation profoundly and shifting the voltage dependence of gating toward more positive potentials (Cherny & DeCoursey, 1999). It was reported recently that the H+ currents that are activated in eosinophils under conditions when NADPH oxidase is functioning have a higher Zn2+ sensitivity than the H+ channels in unstimulated cells and thus are conducted through a different type of H+ channel (Bánfi et al. 1999). Therefore, we examined the effects of ZnCl2 on H+ currents in eosinophils before and after activation with PMA. The effects were qualitatively the same in both conditions. Figure 6 shows H+ currents elicited by identical families of pulses in the same eosinophil in the presence of 1 µM ZnCl2 (Fig. 6A) and after washout with a solution containing EGTA to ensure complete removal of Zn2+ (Fig. 6B). It is evident that the activation of H+ current was slowed profoundly by Zn2+, and that Vthreshold was shifted toward more positive voltages. The cell was then stimulated with PMA, which produced the usual effects on voltage-gated proton channels and turned on -5 pA of Ie. A family of pulses in the presence of 1 µM ZnCl2 is shown in Fig. 6C, and currents during the same family of pulses are shown in Fig. 6D after washout with EGTA-containing solution. The effects of ZnCl2 are quite similar before and after PMA stimulation: profound slowing of activation and a shift of Vthreshold toward more positive voltages. The slowing of activation was so profound that even with longer pulses (16-20 s) in this and other experiments, we could not determine act reliably in the presence of 1 µM ZnCl2. We estimate that the slowing at a given voltage was at least 8- to 10-fold, both in unstimulated and PMA-activated cells. The shift of the gH-V relationship averaged 35-40 mV in five unstimulated eosinophils and three PMA-activated eosinophils. With due consideration of the complexity of the effects of ZnCl2 on voltage-gated proton channels, within the resolution of these measurements we could detect no difference in the type or magnitude of the effects of Zn2+ on voltage-gated proton channels in unstimulated and PMA-activated eosinophils.
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Figure 6. The ZnCl2 sensitivity of H+ current is the same before and after activation with PMA A, a family of currents in an unstimulated eosinophil in the presence of 1 µM ZnCl2 during pulses to 0 mV through +80 mV in 20 mV increments. In contrast with our usual bath solution, this solution had no EGTA. B, the same family of pulses applied after complete removal of ZnCl2 by washout with an EGTA-containing solution. The pulses families in A and B were started 16 and 18 min, respectively, after establishing permeabilized patch recording. The upper calibration bars apply to both A and B. C, approximately 21 min after starting the experiment, PMA was added, and the cell responded in the usual manner: H+ current activation was shifted to a more negative voltage range, | ||
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DISCUSSION |
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H+ current properties in eosinophils in the permeabilized patch configuration
Human eosinophils have the highest reported H+ current density of any native cell, up to hundreds of picoamps per picofarad (Gordienko et al. 1996; Schrenzel et al. 1996). At first glance, the H+ current amplitudes in eosinophils studied here with the permeabilized patch approach appear relatively modest. However, the prodigious H+ current density in whole-cell studies was observed at low pHi (Gordienko et al. 1996; Schrenzel et al. 1996). At pHi 7.0, the IH at the end of a 2 s pulse was 25 pA at +80 mV and 54 pA at +100 mV in the whole-cell configuration (Gordienko et al. 1996). These amplitudes are similar to the representative data shown in Fig. 1 and to the average IH at the end of 8 s pulses to +80 mV of 48 pA in untreated eosinophils studied in permeabilized patch configuration. Because we applied longer pulses, we observed more complete activation at each voltage. Thus, there appears to be general agreement between the amplitudes of H+ currents under both sets of recording conditions. Because previous authors did not report gating kinetics in detail, we cannot compare the kinetics described here directly with those in whole-cell studies of eosinophils. Inspection of published whole-cell data (Gordienko et al. 1996; Schrenzel et al. 1996) suggests that any differences are small.
Like voltage-gated proton currents in other phagocytes (Kapus et al. 1993; DeCoursey & Cherny, 1993), those in eosinophils have slow gating kinetics. The conductance activates upon depolarization with act of several seconds. The recording conditions in the present study are reasonably close to being physiological with respect to pHo, pHi, and having intact cytoplasm. The temperature, to which voltage-gated proton channels are extremely sensitive (Kuno et al. 1997; DeCoursey & Cherny, 1998), is clearly far from physiological. Correcting act measured at +40 mV in unstimulated eosinophils at 20 °C (Fig. 1C) to 37 °C gives ~100 ms, which is a rapid response relative to the ~5 s latency of for H2O2 generation and the half-time of 2.2 s for arachidonic acid release from guinea-pig eosinophils stimulated with leukotriene B4 (Lindsay et al. 1998). Voltage-gated proton channels are thus kinetically competent to be activated as rapidly as the earliest events that occur during a receptor-mediated respiratory burst in eosinophils.
H+ channels in eosinophils were exquisitely sensitive to Zn2+. The effects observed in both unstimulated and PMA-activated cells were similar and were qualitatively like those in all voltage-gated proton channels studied to date (DeCoursey & Cherny, 1994; Cherny & DeCoursey, 1999). There is no evidence of significant simple block, in the sense of state-independent channel occlusion. One micromolar ZnCl2 slowed act by roughly an order of magnitude and shifted the gH-V relationship by 35-40 mV. In the H+ channels of rat alveolar epithelial cells, both of these effects occurred at ~10 µM ZnCl2 at pHo 7 (Cherny & DeCoursey, 1999). H+ channels in eosinophils are thus an order of magnitude more sensitive to Zn2+. This difference reinforces the idea that the epithelial (type e) channel is a different isoform from the phagocyte (type p) channel (DeCoursey, 1998).
Electron currents and O2- release reflect NADPH oxidase activation by PMA
DPI-sensitive inward currents were activated in most human eosinophils upon the initial application of PMA in permeabilized patch conditions. This inward current is a direct reflection of electron efflux across the membrane through the NADPH oxidase enzyme complex and, thus, referred to as electron current, Ie. The following evidence supports this conclusion.
1. The putative Ie is inhibited by DPI.
2. Activation of Ie occurs in permeabilized patch configuration but not whole-cell configuration, which is consistent with it reflecting a diffusible second messenger-mediated response.
3. The kinetics of Ie turn on upon stimulation with PMA are appropriate. The time course was variable, but usually there was a clear delay before the maximal rate of Ie turn on took place. This sigmoidal onset is consistent with the ~1 min delay before the onset of O2- release in time-resolved measurements in human eosinophils at 37 °C (Petreccia et al. 1987), in human neutrophils at 37 °C (Quinn et al. 1993), in pig neutrophils at 22-24 °C (Kapus et al. 1992), and in guinea-pig eosinophils at 25 °C (Lacy et al. 1999).
4. The amplitude of Ie was three times larger in eosinophils than neutrophils, consistent with O2- generation rates reported here (Fig. 3) and in the literature. The peak Ie elicited by PMA was -6.0 pA in eosinophils (Table 1), distinctly larger than -2.3 pA found in neutrophils under similar conditions (DeCoursey et al. 2000). The maximum rate of O2- release measured in eosinophils and neutrophils was 2.8 and 0.87 nmol (105 cells)-1 (5 min)-1, respectively, which can be converted to -9.1 and -2.8 pA of Ie. Both results are in excellent agreement with the roughly 3-fold higher O2- release in human or guinea-pig eosinophils compared with neutrophils stimulated with PMA or other respiratory burst agonists (DeChatelet et al. 1977; Yamashita et al. 1985; Shult et al. 1985; Petreccia et al. 1987; Yagisawa et al. 1996; Someya et al. 1997).
Electron currents were reported previously in human eosinophils dialysed with an artificial solution containing NADPH and other factors (Schrenzel et al. 1998). The peak inward current was -15.6 pA with NADPH in the pipette and -7 pA without NADPH (Schrenzel et al. 1998). The former value is somewhat higher than the present results, but the relevant comparison may be with cells without added NADPH, as our experimental procedure provided no external source of NADPH.
An Ie of -6 pA reflects efflux of 3.75 107 electrons s-1 cell-1. Given an estimate of the total number of functioning NADPH oxidase complexes in each cell, we can calculate the turnover rate of the enzyme. In human neutrophils ~250 000 molecules each of p47phox, p67phox, and rac are translocated to the membrane within 6 min of PMA stimulation (Quinn et al. 1993). It appears that continuous translocation of these components is required for sustained O2- release. If we nevertheless assume that all 250 000 molecules form simultaneously active NADPH oxidase complexes, and scale this number up by a factor of 2-3 to reflect the proportionately higher content of NADPH oxidase components in eosinophils (Segal et al. 1981; Yagisawa et al. 1996; Someya et al. 1997), then each enzyme complex pumps 300-450 electrons s-1 out of the cell. This estimate is in good agreement with the turnover rate of NADPH oxidase in a cell-free system of 240-330 s-1 (Koshkin et al. 1997; Cross et al. 1999). This order of magnitude agreement either confirms the estimate of the number of active NADPH oxidase complexes in each eosinophil or it validates the estimate of the turnover rate of the enzyme, depending on which value one finds more reliable.
Comparison of PMA response of H+ channels with that of neutrophils
The responses of eosinophils to PMA described here are qualitatively similar to those of neutrophils (DeCoursey et al. 2000). In response to PMA, H+ currents activate faster (smaller act), the maximum H+ conductance increases, tail current decay is slowed dramatically, and the gH-V relationship is shifted about 40 mV more negative. There were several quantitative differences between the responses of eosinophils and neutrophils. The PMA-induced changes in H+ currents were generally slightly faster in eosinophils (Table 1). Significantly, the responses in eosinophils were more transient, with the Ie as well as the H+ current parameters often reversing spontaneously, at least partially, within 10-20 min after addition of PMA. In neutrophils, partial spontaneous reversal of the activation of Ie and the slowing of tail in response to PMA occurred infrequently. A more pronounced difference is that in neutrophils we never observed convincing reversal of the PMA-induced increase in IH, the more rapid activation rate, or the hyperpolarizing shift of the gH-V relationship. In contrast, in many eosinophils all of these parameters reversed spontaneously over time. In both neutrophils and eosinophils in which Ie turned off spontaneously, the PMA-induced slowing of tail usually reversed in parallel. It is possible that the PMA enhancement of IH might have reversed eventually in neutrophils, but we saw no evidence of this in some experiments lasting 1 h or more (DeCoursey et al. 2000). The transience of Ie in eosinophils compared with neutrophils is consistent with the more transient time course of PMA-stimulated O2- generation in eosinophils observed here (Fig. 3) and in previous studies (Shult et al. 1985).
All of the changes in H+ channels that occur upon PMA stimulation are in the direction expected to result from cytoplasmic acidification. Because NADPH oxidase activity itself generates intracellular protons, it is reasonable to ask to what extent the changes might be the result of a decrease in pHi rather than modulation of the properties of voltage-gated proton channels. Indeed, Vrev was more negative after PMA treatment indicating that slight cytoplasmic acidification did occur in spite of the NH4+ gradient. The actual change in pHi depends on the balance between two opposing forces. Because there is a direct proportionality between electron translocation across the membrane and protons deposited into the cell by NADPH oxidase, it can be considered that Ie directly indicates the acid load on the cell. Thus Ie of -6 pA reflects a requisite continuous net cytoplasmic acidification equivalent to -6 pA of H+ current (i.e. influx of 3.75 107 H+ s-1 cell-1). Opposing this acid load is the applied NH4+ gradient, whose ability to clamp pHi will be influenced by cell size and pipette geometry. After treatment of eosinophils with PMA, Vrev shifted on average -5.8 mV, corresponding to a decrease in pHi of ~0.1 unit. Because DPI partially reversed the shift, at least part of the pHi shift is attributable to NADPH oxidase-induced acidification. A drop in pHi of 0.1 units will shift the gH-V relationship only -4 mV according to the empirical observation that over a wide range the gH-V relationship changes by 40 mV per unit change in 
pH (Cherny et al. 1995). Thus, at most 10 % of the gH-V shift is ascribable to the lower pHi. The slowing of tail and speeding of act predicted to occur due to a 0.1 unit drop in pHi based on the slopes in Fig. 1C and assuming a -4 mV shift of all kinetic parameters would be 14 and 6 %, respectively. The peak changes in tail and act produced by PMA were 540 and 417 %, respectively (Table 1). Thus, only a tiny fraction of the changes in H+ channel gating kinetics can be ascribed to a loss of control over pHi due to the functioning of NADPH oxidase or other causes. PMA must modulate the properties of H+ channels themselves, rather than changing their microenvironment.
Does PMA activate a new type of H+ channel or modulate existing channels?
Human eosinophils studied in conventional whole-cell configuration express very large voltage-gated proton currents with properties similar to those in other phagocytes (Gordienko et al. 1996; Schrenzel et al. 1996). However, in human eosinophils studied in whole-cell configuration with NADPH and other factors included in the pipette to elicit NADPH oxidase function, the properties of voltage-gated proton channels were markedly different (Bánfi et al. 1999). The gH activated in a much more negative voltage range, with the result that inward H+ currents were activated negative to EH at some pH. In addition, tail was quite slow. Bánfi et al. (1999) hypothesized that a novel type of voltage-gated proton channel was activated when the NADPH oxidase was working. This H+ channel (type x) was postulated to be distinct from the type p H+ channels observed in conventional whole-cell technique. In unstimulated eosinophils in permeabilized patch configuration proton currents exhibit type p properties like those in whole-cell studies (Gordienko et al. 1996; Schrenzel et al. 1996). After PMA activation the H+ currents exhibit clear type x properties, including activation of inward currents and very slow tail.
In neutrophils, we concluded that a single population of H+ channels was present and that PMA greatly alters the properties of these channels but does not induce a new type of channel (DeCoursey et al. 2000). The gH increased in direct proportion to the resting gH, consistent with modulation rather than induction of an independent conductance. Tail current decay was monoexponential before, during and after the slowing by PMA both in neutrophils and in many eosinophils. Similarly, the gH-V relationship did not exhibit evidence of two components. The strongest evidence for the existence of two distinct types of channels was that type x currents seemed to be 20-fold more sensitive to Zn2+ inhibition than type p currents (Bánfi et al. 1999). In contrast, in the present study, the inhibition of H+ currents by Zn2+ in human eosinophils was similar in unstimulated and PMA-activated cells. In the Appendix, we show that the apparent differences between our conclusions and those of Bánfi et al. (1999) reflect differences in the way that Zn2+ effects were assessed. The similarity of Zn2+ sensitivity of H+ currents in eosinophils before and after PMA activation provides further support for the idea that a single population of H+ channels exists. PMA strongly modulates the properties of these channels, greatly enhancing their probability of opening.
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APPENDIX |
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We interpret the effects of PMA on proton channels in human eosinophils as modulation of the properties of channels that are present in the surface membrane of resting cells. In the present study, we see evidence of only one population of channels, and by our criteria, the Zn2+ sensitivity was the same in unstimulated and PMA-activated eosinophils. In contrast, Bánfi et al. (1999) proposed that eosinophils express two distinct varieties of voltage-gated proton channels. A 'high-threshold' channel that opens at voltages positive to EH is present in the surface membrane of unstimulated cells. A 'low-threshold' channel is absent from resting cells but appears when NADPH oxidase is active. This channel is unique among H+ channels in activating negative to EH under some conditions, resulting in inward current. The 'low-threshold' channel was reported to be 20-fold more sensitive to inhibition by Zn2+ (Bánfi et al. 1999). This conclusion was based on comparison of concentration- response relations at two voltages: at +20 mV where (under their conditions) there was inward H+ current attributed to the 'low-threshold' conductance, and at +60 mV where 'high-threshold' channels would also be activated. The concentration-response curve was shifted toward higher [Zn2+] at the more positive test voltage. In this Appendix we show that these results are qualitatively consistent with the existence of a single population of voltage-gated proton channels. In fact, the apparent Zn2+ potency as measured by Bánfi et al. (1999) must depend on test potential to the extent that its effects are due to its ability to shift the voltage dependence of H+ channel gating.
Zn2+ and other divalent metals have several effects on all voltage-gated proton channels that have been studied, and have similar effects on many other types of ion channels. The voltage-activation curve is shifted to more positive voltages, the activation of current during a depolarizing pulse is slowed, and maximal gH may be decreased. We report here that Zn2+ exhibits these effects, in particular the depolarizing voltage shift, in human eosinophils both before and after activation by PMA. It is not easy to distinguish among these effects; some slowing of act is expected from the voltage shift of gating. For the present purpose, we test the simple assumption that the entire effect of Zn2+ is due to its voltage-shifting ability, which results from Zn2+ binding to a site at the external side of the channel (Cherny & DeCoursey, 1999) with affinity KM = 10-7 M (set arbitrarily for the purpose of illustration). We describe the gH-V relationship in the absence of Zn2+ as a Boltzmann distribution with midpoint at 0 mV and a slope factor of 10 mV. As discussed previously (p. 832 of Cherny & DeCoursey, 1999), given simple assumptions, the voltage shift actually produced by Zn2+ can be described empirically from the slope factor of the gH-V relationship and the KM of Zn2+. Figure 7A shows the predicted voltage shift as a function of Zn2+ concentration; the inset illustrates a voltage shift at one arbitrary concentration of Zn2+.
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Figure 7. The depolarizing shift of voltage-dependent gating by ZnCl2 results in test-pulse dependence of apparent Zn2+ sensitivity A, the dependence of the voltage shift of the gH-V relationship on ZnCl2 concentration is illustrated for a semi-empirical model of Zn2+ effects on voltage-gated proton channels (Cherny & DeCoursey, 1999). The numerical value of the shift was derived by assuming that the channel cannot open while Zn2+ is bound to it. The open probability of the channel in the absence of Zn2+ is given by a simple Boltzmann relationship: Popen = (1 + exp((V - V1/2)/k))-1 where the mid-point potential V1/2 was set arbitrarily at 0 mV and the slope factor k is -10 mV, a value typical for voltage-gated proton channels in many cells (DeCoursey & Cherny, 1994). By definition, if k = -10 mV then Popen is reduced e-fold in 10 mV. Hence, if Zn2+ reduced Popen e-fold this would shift the gH-V relationship by 10 mV (Cherny & DeCoursey, 1999). The curve is drawn according to: Vshift = ln(1 - PZn)-1 | ||
Ion channel block is often quantified as the ratio of the current in the presence of blocker to that in its absence. This procedure was used by Bánfi et al. (1999). The implicit assumption is that the blocker simply occludes the channel, with no dependence on voltage or the gating state of the channel, and conversely, the blocker does not alter the gating kinetics or voltage dependence of the channel. It is abundantly clear that Zn2+ has profound effects on both the gating kinetics and voltage dependence of voltage-gated proton channels (Cherny & DeCoursey, 1999). In fact, 'block' in the sense of simple channel occlusion is at most a minor part of the actions of Zn2+. The inset in Fig. 7A illustrates how a simple voltage shift affects the current measured at different voltages, V1 and V2. The fractional decrease in H+ current measured at the end of the pulse is much greater at test voltage V1, as can also be seen from the intersection of the lines in the schematic gH-V relationships. Thus, the apparent affinity of Zn2+ for the H+ channel will depend on the test voltage.
Figure 7B illustrates the calculated concentration- response relationships that result when Zn2+ effects are evaluated as the ratio of current in the presence and absence of Zn2+. The curves calculated for different test voltages resemble experimental data that were analysed in this way (Bernheim et al. 1993; Gordienko et al. 1996; Bánfi et al. 1999). Except at negative voltages where the open probability of the channel is very low, the apparent potency of Zn2+ is strongly dependent on the test voltage, with the calculated apparent Ki increasing about 20-fold for a test voltage 40 mV more positive. Bánfi et al. (1999) reported that the Ki of Zn2+ was 20-fold higher when the test voltage was +60 mV compared with +20 mV. Thus, rather than comprising evidence of two types of channels, the Zn2+ concentration-response data of Bánfi et al. (1999) are entirely consistent with the existence of a single population of proton channels.
The dependence of the reduction of H+ current by Zn2+ on test voltage gives the impression that 'block' is voltage dependent. However, genuine voltage-dependent block differs in two significant ways from the phenomenon in Fig. 7B. First, voltage-dependent block suggests that Zn2+ acts at a site within the membrane potential field. There is no evidence that external Zn2+ receptor of voltage-gated proton channels is within the membrane potential field, as discussed at length elsewhere (Cherny & DeCoursey, 1999). If the conduction pathway through voltage-gated proton channels is a hydrogen-bonded chain including side groups of amino acids, as has been suspected for some time (DeCoursey & Cherny, 1994), then Zn2+ could not permeate. Second, voltage-dependent block implies that the time Zn2+ occupies its receptor depends on voltage. There is no evidence that this is the case, and Fig. 7B illustrates that existing data are well described by assuming that Zn2+ occupies its receptor independently of membrane potential.
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