INTRODUCTION

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Voltage-gated H⁺ channels were described first in snail neurons (Thomas and Meech, 1982; Byerly et al., 1984), and more recently have been found in mammalian cells (DeCoursey, 1991). A recent burst of interest in this channel has resulted from its proposed role in extruding protons from neutrophils and other phagocytes. During the "respiratory burst," NADPH oxidase secretes superoxide anion to kill bacteria and simultaneously releases protons into the cytoplasm. Henderson and colleagues (1987, 1988a, 1988b) deduced that electrogenic H⁺ efflux provided the necessary charge compensation, on the basis of pH and membrane potential changes in human neutrophils. The presence of voltage- and pH-activated H⁺-selective channels in human neutrophils was subsequently confirmed by voltage clamp (DeCoursey and Cherny, 1993). Despite increasing interest in this channel, its molecular identity has not been established and numerous questions about the properties and physiological regulation of this channel remain unanswered.

Here we explore the effects of temperature on two fundamental properties of voltage-activated H⁺ channels: pH-dependent gating and H⁺ permeation. Interpreting the results requires distinguishing the effects of temperature on the voltage- and pH-dependent gating mechanism from those on the conductance of the open channel. A recent suggestion that H⁺ currents in murine mast cells have a greater temperature sensitivity than other H⁺ channels (Kuno et al., 1997) led us to examine the temperature dependence in several mammalian cells and cell lines. In addition, it is now clear that the properties of H⁺ channels differ in different cells. We studied H⁺ currents in rat alveolar epithelial cells, rat macrophages, human neutrophils, human monocyte THP-1 cells, human promyelocyte HL-60 cells, and mouse microglial BV-2 cells. These cells include both type e (epithelial) and p (phagocyte) H⁺ channel varieties (DeCoursey, 1998) and 6 of the 17 mammalian cells or cell lines and 3 of 5 mammalian species in which H⁺ currents have been reported. We find similarly high temperature sensitivity in all mammalian cells.

Voltage-gated H⁺ channels are extremely selective for H⁺ (and deuterium), with no detectable permeability to other cations (Barish and Baud, 1984; Byerly et al., 1984; DeCoursey, 1991; Bernheim et al., 1993; Kapus et al., 1993; Demaurex et al., 1993; DeCoursey and Cherny, 1994a, 1994b, 1997; Gu and Sackin, 1995; Gordienko et al., 1996; Kuno et al., 1997). Although the macroscopic conductance increases at lower pH_i, the increase is only 1.7-fold/U decrease in pH_i when measured in inside-out patches (DeCoursey and Cherny, 1995, 1996a), far less than the 10-fold increase expected if the conductance were proportional to the permeant ion concentration, [H⁺]_i. In contrast, the H⁺ conductance of gramicidin (Akeson and Deamer, 1991; Cukierman et al., 1997) and several other H⁺ permeable channels (reviewed by DeCoursey and Cherny, 1994b) increases in direct proportion to [H⁺] up to pH ~0, and then saturates. Thus, the conductance of the voltage-gated H⁺ channel appears to be nearly saturated at pH 7. Because relatively small changes in g_H are seen when either intracellular or extracellular buffer concentrations were varied 100-fold (DeCoursey and Cherny, 1996b), neither buffer diffusion nor direct proton transfer from buffer to channel can be rate determining steps in conduction. The ratio of H⁺ to D⁺ current was 1.9 at 20°C (DeCoursey and Cherny, 1997), much larger than 1.41-1.52 for the ratio of H⁺ to D⁺ conductivities in bulk solution at 20°C (Lewis and Doody, 1933; Roberts and Northey, 1974). Taken together, these studies suggest that the rate-determining step in H⁺ permeation occurs in the pore rather than in the diffusional approach of either protonated buffer or H₃O⁺. The activation energy (E_a)¹ reported here for H⁺ permeation is large enough to rule out conclusively the possibility that diffusion to the channel is rate determining. That the E_a is as high as for hydrolysis leads to renewed consideration of this mechanism (Kasianowicz et al., 1987) as a possible source for a fraction of the protons that carry current through these channels.

A quintessential feature of all voltage-gated H⁺ channels is the striking dependence of their voltage-gating mechanism on pH_o and pH_i. This interaction was examined systematically in alveolar epithelial cells, where the voltage-activation curve was found to shift 40 mV/U increase in the pH gradient, pH = pH_o  pH_i (Cherny et al., 1995). The threshold voltage at which the g_H is first detectably activated can be predicted from:

(1)

where V₀ was typically 20 mV (Cherny et al., 1995), or:

(2)

where V_rev is the observed reversal potential (DeCoursey and Cherny, 1997). The importance of the regulation of gating by pH is that the g_H is activated only when there is an outward pH, thus the channel evidently functions to extrude H⁺ from the cell. We have proposed that the regulation of gating by pH is mediated by internal and external protonation sites, which are accessible only to one side of the membrane at a time, and whose accessibility is governed by a conformational change in the channel molecule that can occur only when the sites are deprotonated (Cherny et al., 1995). The effects of temperature on gating kinetics further elucidate the gating mechanism. The surprising similarity of the Q₁₀ for activation and deactivation suggests that the same process is rate determining for both opening and closing of H⁺ channels.

	MATERIALS AND METHODS

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Cells

Human neutrophils. Neutrophils were isolated from normal human blood by density gradient centrifugation (Schmeichel and Thomas, 1987), and kept on ice in nominally divalent-free buffer. Immediately before recording, neutrophils were transferred to the glass recording chamber and superfused with Ringer's solution (see Solutions, below). In some experiments, fresh blood from the authors was studied without purification, and neutrophils were identified visually by their size (~8 µm diameter) and spherical, granular appearance, as described previously (DeCoursey and Cherny, 1993).

THP-1 cells. THP-1 cells were obtained from American Type Culture Collection (Rockville, MD). Cells were cultured in suspension in RPMI medium supplemented with 0.29 mg/ml glutamine, 10% fetal bovine serum (Gibco Laboratories, Grand Island, NY), 100 U/ml of penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml Fungizone (Amphotericin B; Gibco Laboratories). Cells were incubated at 37°C in a humidified atmosphere of 5% CO₂ in air. Every 2-3 d, about half of the media was replaced with fresh media, and once per week the cells were removed, centrifuged at 1,800 rpm for 10 min at 4°C in an RT6000 refrigerated centrifuge with an H1000B rotor (both from Sorvall, Newtown, CT). The cell pellet was resuspended in fresh media at 1-2 × 10⁶ cells/ml. THP-1 cells are nonadherent. With maintained weak positive pressure, the pipette was placed on or near a cell, and then suction was initiated.

BV-2 cells. BV-2 cells were a gift from Claudia Eder (University of California at Irvine, Irvine, CA). The cells were maintained in DMEM with 10% FCS and 1% L-glutamine.

HL-60 cells. HL-60 cells were obtained from American Type Culture Collection. They were grown in RPMI 1640 media containing 20% FCS. Some cells were studied after treatment with 1% DMSO for 7 d to induce differentiation into granulocytes.

Rat alveolar epithelial cells. Type II alveolar epithelial cells were isolated from adult male Sprague-Dawley rats using enzyme digestion, lectin agglutination, and differential adherence, as described in detail elsewhere (DeCoursey et al., 1988; DeCoursey, 1990), with the exception that we now use elastase without trypsin to dissociate the cells. Some earlier experiments on cells isolated with trypsin and elastase are included here. Before invasive procedures were initiated, the rats were anesthetized deeply using sodium pentobarbital. In brief, the lungs were lavaged to remove macrophages, elastase was instilled, and then the tissue was minced and forced through fine gauze. Lectin agglutination and differential adherence further removed contaminating cell types. The preparation at first includes mainly type II alveolar epithelial cells, but after several days in culture the properties of the cells are more like type I cells. No changes in the properties of H⁺ currents have been observed during this differentiation process. H⁺ currents were studied in approximately spherical cells up to several weeks after isolation.

Rat macrophages. Rat macrophages were obtained by lavage during the isolation of type II alveolar epithelial cells. They were studied <1 d after removal from the rat.

Solutions

Most solutions (both external and internal) contained 1 mM EGTA, 2 mM MgCl₂, and 100 mM buffer, with tetramethylammonium methanesulfonate (TMAMeSO₃) added to bring the osmolarity to ~300 mosM, and titrated to the desired pH with tetramethylammonium hydroxide (TMAOH) or methanesulfonic acid (solutions using bis-Tris as a buffer). The pH 7 and 8 solutions contained 3 mM CaCl₂ instead of MgCl₂. A stock solution of TMAMeSO₃ was made by neutralizing TMAOH with methanesulfonic acid. Buffers (Sigma Chemical Co., St. Louis, MO), which were used near their pK in the following solutions, were: pH 5.5-6.0 Mes; pH 6.5 bis-Tris (bis[2-hydroxyethyl]amino-tris[hydroxymethyl]methane); pH 7.0 Bes (N,N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid); pH 7.5 HEPES; pH 8.0 Tricine (N-tris[hydroxymethyl]methylglycine). In a few experiments (done 5-6 yr ago), TEA⁺ replaced TMA⁺, and 20 mM buffer was used. Whether TEA⁺ is inert with respect to H⁺ channels is uncertain, but the temperature dependence of the currents appeared consistent with other data using TMA⁺. In a few other experiments, N-methyl-D-glucamine was used as an impermeant cation instead of TMA⁺. The initial bath solution was usually Ringer's solution containing (mM): 160 NaCl, 4.5 KCl, 2 CaCl₂, 1 MgCl₂, 5 HEPES, pH 7.4.

Electrophysiology

Conventional whole-cell, cell-attached, or excised inside-out patch configurations were used. Inside-out patches were generally formed by lifting the pipette into the air briefly. Micropipettes were pulled in several stages using a Flaming Brown automatic pipette puller (Sutter Instruments, Co., San Rafael, CA) from EG-6 glass (Garner Glass Co., Claremont, CA), coated with Sylgard 184 (Dow Corning Corp., Midland, MI), and heat polished to a tip resistance ranging typically from 3 to 10 M. Electrical contact with the pipette solution was achieved by a thin sintered Ag-AgCl pellet (In Vivo Metric Systems, Healdsburg, CA) attached to a silver wire covered by a Teflon tube. A reference electrode made from a Ag-AgCl pellet was connected to the bath through an agar bridge made with Ringer's solution. The current signal from the patch clamp (List Electronic, Darmstadt, Germany) was recorded and analyzed using an Indec Laboratory Data Acquisition and Display System (Indec Corp., Sunnyvale, CA). Data acquisition and analysis programs were written in BASIC23 or FORTRAN. Seals were formed with Ringer's solution in the bath, and the zero current potential established after the pipette was in contact with the cell. Inside-out patches were formed by lifting the pipette into the air briefly.

Pulse duration. Pulse duration was adjusted at different temperatures with the intent of balancing two opposing factors. Longer pulses tend to provide a better estimate of both gating kinetics and steady state current amplitudes. However, the longer the pulse, the greater the increase in pH_i caused by H⁺ efflux- mediated depletion of protonated buffer from the cell. Depletion directly distorts the H⁺ current waveform and also necessitates long interpulse intervals to allow pH_i to recover. Most whole-cell measurements above 30-35°C were plagued by signs of changes in bulk pH_i due to the massive H⁺ extrusion during voltage pulses, even though we tried to use relatively small and brief depolarizations to avoid this complication. We generally stopped increasing the temperature when pronounced droop of the H⁺ current occurred.

Temperature Control

Bath temperature was controlled by Peltier devices in a feedback arrangement, and was monitored by a resistance temperature detector (RTD) element (Omega Scientific, Stamford, CT) placed in the bath near the cell. Bath temperature was recorded at the end of the pulse just before writing to disk, and was stored with each current record. The maximum rate of change of bath temperature was ~0.1°C/s. During temperature changes, the temperature often changed during long pulses. We usually stayed for several minutes at temperatures in intervals of 5-10°C to fix the behavior more accurately. Before lowering the bath temperature, we lifted the cell (via the pipette) because otherwise thermal contraction of the copper housing supporting the bath lifted the chamber enough to smash the pipette tip.

Temperature effects on buffer pK_a. The pK_a of most buffers decreases with increased temperature by 0.01-0.02 U/°C. When we change the temperature, the pK_a of the buffers used to establish the pH of internal and external solutions will change, and consequently H⁺ will be released or bound by buffer. When the same buffer, or buffers with similar temperature dependence, are used in the bath and pipette solutions, temperature will not affect the pH gradient, pH, but will change the absolute pH. However, when buffers with different temperature dependences are used in the bath and pipette solutions, pH (as well as the absolute pH) will change. Most experiments were done with Mes or bis-Tris in the pipette solution, both of which have weaker temperature dependence than the most frequently used extracellular buffers (Bes, less often HEPES, Tricine, or others). Consequently, pH will decrease at higher temperatures, and E_H will generally change less at higher temperatures and in some situations actually decrease (because changes in pH and T in the Nernst equation tend to cancel each other). Over a temperature range spanning 30°C for the buffers used, the largest change in absolute pH is 0.42 U (for Tricine), and the largest net change in pH is ~0.26 U for pH 8.0//6.5 (Tricine//bis-Tris). Most measurements were done at pH 7.0//5.5 (Bes//Mes), where pH changes 0.04 U/10°C. All solutions are described according to their nominal pH when titrated at room temperature (20-24°C).

Conventions

We refer to pH in the format pH_o//pH_i. In the inside-out patch configuration, the solution in the pipette sets pH_o, defined as the pH of the solution bathing the original extracellular surface of the membrane, and the bath solution sets pH_i. Currents and voltages are presented in the normal sense; that is, upward currents represent current flowing outward through the membrane from the original intracellular surface, and potentials are expressed by defining as 0 mV the original bath solution. 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 delay (as described in DeCoursey and Cherny, 1995):

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 t_delay is the delay. The H⁺ current amplitude (I_H) is defined as (I₀  I). No other time-dependent conductances were observed consistently under the ionic conditions employed. The tail current time constant, _tail, was obtained by fitting the current with a single exponential:

where I₀ is the amplitude of the decaying part of the tail current.

Calculation of Q₁₀ or Arrhenius activation energies. The relative change in a parameter for a 10°C change in temperature, the Q₁₀, was calculated by:

(3)

(4)

1

1

	RESULTS

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The amplitude and kinetics of voltage-gated H⁺ currents are strongly temperature dependent. For example, evaporative heat loss from the solution in the recording chamber lowers the temperature by ~1°C, producing a noticeable change in the H⁺ currents. Fig. 1 illustrates families of H⁺ currents recorded in a human neutrophil at 11, 20, and 36°C (note the different time bases). Increasing the temperature by 25°C increased the H⁺ current amplitude >7-fold and the rate of activation (turn on) of the current with depolarization >20-fold. At all temperatures, H⁺ currents activated during depolarizing pulses with a sigmoidal time course, suggesting that the channel passes through more than one closed state before opening (DeCoursey and Cherny, 1994b; Cherny et al., 1995). In general, the behavior of H⁺ currents appeared to be fairly consistent at all temperatures; that is, after scaling the amplitude and time scales, there were no obvious changes in the characteristic properties or appearance of the current waveforms.

View larger version (11K): [in this window] [in a new window]   Fig. 1.   Families of H+ currents in a human neutrophil at 11, 20, and 36°C. From Vhold = 60 mV, pulses were applied in 20-mV increments from 40 to +40 mV. Bath solution was TMAMeSO3 at pHo 7.0; the pipette contained NMGMeSO3 at pHi 5.5. The current calibration bar applies to all panels; note the change in time calibration.

Fig. 2 illustrates tail currents and V_rev measurements in the same neutrophil as in Fig. 1. A depolarizing prepulse opened many H⁺ channels, and then the membrane was repolarized to various potentials. The time course of current decay reflects the progressive closing of channels at each potential. The tail current decay was well fitted by a single exponential, giving _tail, the time constant. The rate of channel closing (1/_tail) is evidently highly temperature sensitive (note the change in time base), as was the rate of channel opening in Fig. 1. When _tail was measured over a wide voltage range at different temperatures (Fig. 2 C), the _tail-V relationship appeared to scale uniformly at all potentials. Our model (Cherny et al., 1995) predicts a 5% steeper slope of the _tail-V relationship at higher temperatures (V.S. Markin, personal communication). However, the measurement is not sufficiently accurate to detect this subtle a change. To a first approximation, the Q₁₀ of _tail is independent of voltage.

View larger version (9K): [in this window] [in a new window]   Fig. 2.   Tail currents at 20.5 (A) and 36.5°C (B) in the same human neutrophil as in Fig. 1 (also at pH 7.0//5.5). The temperature varied several tenths of a degree from pulse to pulse and we have rounded to the nearest half degree. From Vhold = 60 mV, a prepulse to +30 (A) or +20 (B) mV was followed by test pulses to the indicated voltages, shown in 20-mV increments. (C) Voltage dependence of tail at 20.5 and 36.5°C in the same experiment as A and B. The slopes are similar, 39 and 44 mV/e-fold change in tail, respectively, giving an average Q10 of 8.5 in this cell.

The reversal potential V_rev was determined from tail current measurements like those in Fig. 2, A and B. In the illustrated neutrophil, V_rev was close to 80 mV at both 20.5 and at 36.5°C. The Nernst potential for H⁺, E_H, at nominally pH_o 7.0//5.5, is 87 mV at 20.5°C and 88 mV at 36.5°C after correction for shifts in the true pH due to the temperature dependence of the pK_a of the buffers used (see MATERIALS AND METHODS). In other cells, V_rev generally was unchanged or somewhat more positive at lower temperatures. The actual value obtained for V_rev is sensitive to depletion of protonated buffer from the cell by previous pulses, including the prepulse used in standard tail current measurements. Within the reliability of the measurement, temperature does not appear to alter V_rev beyond the small change in E_H predicted by the Nernst equation.

The steady state voltage dependence of the g_H (the activation curve) was generally similar at all temperatures. This result is important partially for technical reasons. The comparison of H⁺ current kinetics at a fixed test potential at different temperatures would become less valid if the P_open were different. A hyperpolarizing shift in P_open at high temperature would tend to artificially enhance the temperature dependence of the activation time constant, _act, and perhaps reduce the temperature dependence of _tail. The voltage dependence of channel opening is not easy to evaluate quantitatively because the chord conductance (g_H) often does not saturate, the g_H-V relationship is poorly described by a Boltzmann function, whole-cell currents are susceptible to depletion effects, and for other reasons discussed at more length elsewhere (DeCoursey, 1991; DeCoursey and Cherny, 1994a, 1994b; Cherny et al., 1995). Therefore, several approaches were taken (not all are illustrated). In Fig. 3 A, g_H was determined in an inside-out patch from an alveolar epithelial cell, from the amplitude of an exponential fit to the rising phase of H⁺ currents. Using the fitted amplitude corrects data in which activation did not achieve steady state during the pulse. In this experiment, activation appeared to occur at more negative potentials (by ~10-15 mV) at higher temperatures. However, both the tremendous increase in gating kinetics, as well as the increase in H⁺ current amplitude, will tend to give this impression, even if there were no true shift. Small changes in pH and E_H due to temperature effects on buffers predict net shifts of a few millivolts in the depolarizing direction at higher temperatures (see Fig. 3, legend). We also looked for changes in V_threshold, the threshold potential at which time-dependent outward current was first detectable. In some experiments, careful examination revealed little or no shift, whereas in other experiments shifts of 5-10 mV occurred, with activation usually occurring at more negative voltages at higher temperatures. The impression gained from these attempts was that any shift in the voltage-activation curve was smaller than could be demonstrated convincingly. We cannot distinguish whether increasing the temperature produced a small (~10 mV) hyperpolarizing shift or no effect. Likewise, we cannot resolve whether high temperature might have steepened the g_H-V relationship somewhat as predicted by our model (Cherny et al., 1995).

View larger version (13K): [in this window] [in a new window]   Fig. 3.   The voltage dependence of H+ current activation does not change much with temperature. (A) Chord conductance-voltage relationship in an inside-out patch from a rat alveolar epithelial cell, at the indicated temperatures, at pHo 7.5 (pipette)//pHi 6.5 (bath). The order of measurement was 20, 10, 21, 30, and 40°C. At 40°C, gH at +20 mV was 1.24 nS (not plotted). Because of the difference in temperature dependence of the buffers used (the pKa of bis-Tris decreases 0.008/°C and HEPES 0.014/°C), the pH gradient across the cell membrane, pH (pHo pHi), will decrease by 0.06 U every 10°C. Assuming that the gH-V relationship shifts 40 mV/U increase in pH (Eq. 1), this effect would give rise to a depolarizing shift of 4.8 mV (or 2.3 mV if the gH-V relation is set by Vrev according to Eq. 2) as the temperature is increased from 10 to 30°C. (B) Voltage dependence of act at various temperatures in the same patch. The lines show the slopes by linear regression, which range from 46 to 64 mV/e-fold change in act and did not change consistently with temperature.

The time course of H⁺ currents, at least for moderate depolarizing pulses, was reasonably well described by a single exponential rise after a delay. Fig. 3 B illustrates the voltage dependence of the activation time constant, _act, measured at several temperatures in the inside-out patch shown in Fig. 3 A. As was found for _tail, the Q₁₀ appeared to be independent of the voltage at which the measurement was made. Our model (Cherny et al., 1995) predicts some temperature dependence of the slope of the _act-V relationship. However, within the accuracy of the measurement, both kinetic parameters appeared simply to scale uniformly with temperature at all voltages. Similar results were obtained in whole-cell experiments.

To evaluate H⁺ channel gating and conductance over a wide range of temperature, we used a moderate depolarizing test pulse, followed by repolarization either to V_hold or to another voltage at which the tail current decay was resolvable. This pulse protocol permitted obtaining all four parameters nearly simultaneously: _act, the delay time, _tail, and I_H. As the temperature was varied, the pulse durations were adjusted to resolve the kinetic parameters. Low temperatures required very long pulses (up to 80 s), with the result that the temperature sometimes changed significantly during the pulse. Some hysteresis in the data is to be expected in this situation. Fig. 4 illustrates the temperature dependence of the parameters studied in a rat alveolar epithelial cell. All four parameters have roughly linear temperature dependence (in a semi-log plot), and thus each can be described by a single Q₁₀. The same data are plotted in Fig. 4 B in a conventional Arrhenius plot in which the slope gives the E_a.

View larger version (18K): [in this window] [in a new window]   Fig. 4.   (A) Temperature dependence of the amplitude and gating kinetics of H+ currents in an alveolar epithelial cell at pH 7.0//6.5. Pulses were applied from Vhold = 20 to +50 mV, and then to 0 mV. The durations of these pulses were varied from 0.5-25 s according to the temperature range. The rising currents at +50 mV were fitted by a delay () followed by a single exponential to obtain act () and the extrapolated H+ current amplitude, IH (). Upon repolarization to 0 mV, the tail current was fitted by a single exponential to obtain tail (). The delay was the least well defined parameter. The slopes, determined by linear regression, correspond with Q10 values of 11.9 for act, 7.09 for the delay, 7.51 for tail, and 2.16 for IH, as illustrated by the lines. (B) Arrhenius plots of the data in A. Plotted against 1,000/temperature (°K1) are the natural logarithm of 1/delay (), 1/act (), 1/tail (), and IH (). The slopes, determined by linear regression, correspond with Ea values of 41.7 kcal/mol for act, 33.0 kcal/mol for the delay, 33.3 kcal/mol for tail, and 13.0 kcal/mol for IH, as illustrated by the lines.

Comparison among Different Cells and Mammalian Species

Fig. 5 illustrates the temperature dependence of the four parameters studied in several types of cells. Several features are immediately apparent. First, the three kinetic parameters, _act, delay, and _tail are all highly temperature sensitive. A second result that is somewhat unusual in temperature studies on ion channels is that there are no obvious changes in the slopes of these curves. The data were well described by a straight line on the graphs, giving a single Q₁₀ value over the entire temperature range. Q₁₀ values from experiments like these are summarized in Table I. The third result evident in both Fig. 5 and Table I is that the three kinetic parameters have nearly the same Q₁₀. The similarity of Q₁₀ values for the three kinetic parameters explains why the general appearance of the currents appeared to simply scale with temperature (compare Figs. 1 and 2).

View larger version (46K): [in this window] [in a new window]   Fig. 5.   Temperature dependence of H+ current parameters in several cell types. (A) Temperature dependence of the amplitude and gating kinetics of H+ currents in a human neutrophil at pHo 7.0//pHi 5.5. From Vhold = 60 mV, pulses were applied to +20 mV with a duration 0.4-8.0 s according to the temperature range. The rising current at +20 mV was fitted by a delay (), followed by a single exponential to obtain act () and IH (). The tail current upon repolarization to 60 mV was fitted by a single exponential to obtain tail (). The delay was the least well defined parameter. The slopes, determined by linear regression, correspond with Q10 values of 4.46 for act, 6.82 for the delay, 4.53 for tail, and 2.33 for IH, as illustrated by the lines. There was noticeable hysteresis in this experiment, as the temperature was first increased from room temperature (19°C) to 37°C, decreased to 11°C, and then returned to room temperature. As a result of this hysteresis, there is a certain amount of arbitrariness in the evaluation of these data. (B) THP-1 cell at pHo 7.0//pHi 5.5. Pulses were applied from Vhold = 60 to +50 mV, and then 30 mV. (C) BV-2 cell at pHo 7.0//pHi 5.5. Pulses were applied from Vhold = 80 to 30 mV, and then 80 mV. (D) Inside-out patch from an alveolar epithelial cell at pHo 7.5//pHi 6.5. Pulses were applied from Vhold = 60 to +20 mV, and then 40 mV. Because the currents in patches were much smaller than whole-cell currents, the kinetic parameters are less well determined, but IH is probably more reliable because depletion effects are minimized (see text).

The H⁺ current amplitude increased substantially with temperature. However, its measurement was complicated by several factors. At low temperature, activation was very slow, and the pulses were sometimes not long enough to reach steady state. We extrapolated the fitted exponential rise to obtain I_H. At high temperatures, the currents were quite large and we sometimes observed decay of outward currents above ~30°C. H⁺ channels do not inactivate (DeCoursey and Cherny, 1994b), and this decay is almost certainly the result of increased pH_i due to the large efflux of H⁺ comprising current flow (see DISCUSSION). Other factors that could artifactually decrease I_H at high temperature include spontaneous clogging of the pipette tip and movement of the cell relative to the pipette due to thermal expansion of the copper support for the chamber (used to transfer heat to the preparation). Because many of these complications can be avoided or minimized using the inside-out patch configuration, we considered this approach to provide the most reliable estimate of the temperature dependence of I_H.

Inside-Out Patch Experiments

Several measurements were made in inside-out patches excised from rat alveolar epithelial cells. Although the smaller current amplitudes compared with whole-cell measurements made the extraction of kinetic parameters less precise, the problems associated with pH_i changes due to depletion of protonated buffer by H⁺ currents are greatly reduced. Fig. 6 illustrates families of H⁺ currents at several temperatures in an inside-out patch. The parameters that could be extracted are plotted in Fig. 5 D, as was done for whole cell data. At higher temperatures, the time-independent "leak" current appears to increase. Because the I_H amplitude is obtained by fitting the current to a delay plus a single exponential, it reflects only the time-dependent component of outward current. If some part of the initial jump in outward current at higher temperature reflects H⁺ current, for example if the decay of capacity current obscures the initial rise in I_H at higher temperatures where activation becomes rapid, then the Q₁₀ of I_H will be underestimated by this procedure. We could not resolve tail currents well enough to estimate _tail reliably over a large temperature range in most patches. The values of _act, summarized in Table I, are generally similar to those from whole-cell experiments. Notably different is I_H. The H⁺ current amplitude in patches continued to increase at higher temperatures (instead of saturating above ~30°C). The Q₁₀ extracted from patches even in the higher temperature range (>20°C, Table I) was larger than for whole-cell experiments. Because the use of excised patches minimizes depletion problems, we believe that these data are more reliable than those obtained in whole-cell experiments.

View larger version (8K): [in this window] [in a new window]   Fig. 6.   H+ current families at 10 (A), 20 (B), and 30°C (C) in an inside-out patch from an alveolar epithelial cell at pHo 7.5// pHi 6.5. In each family, Vhold was 60 mV, and pulses to 20, 0, and +20 mV are illustrated. Note the changes in time calibration. The same current calibration applies to all parts.

The I_H data from patches exhibited clear nonlinearity of the type observed to a lesser extent in whole cell I_H data. In Fig. 7, I_H data from three patches with different current amplitudes are plotted. The Q₁₀ is 2-3 at 20-30°C, but much larger (4-6) at temperatures below 20°C. The curvature is in the direction that would be attenuated in an Arrhenius plot; however, the Arrhenius plots in Fig. 7 B are still clearly nonlinear, with E_a ~15 kcal/mol at high temperatures and ~30 kcal/mol at low temperatures. Activation energies clearly vary depending on the temperature range studied. In Tables I we list values from above and below 20°C.

View larger version (15K): [in this window] [in a new window]   Fig. 7.   Nonexponential temperature dependence of IH in three inside-out patches from alveolar epithelial cells. (A) IH in three patches (one is also plotted in Figs. 5 D and 6). Because the plots are not linear, we arbitrarily fitted the data at <21 (, ), 21-35 (, ), and >35°C (). Lines determined by linear regression are drawn through the points and Q10 values are given near the relevant sections of data. (B) The same data in an Arrhenius plot. Although the curvature is slightly less than in A, the plot clearly is steeper at low temperatures. Dots inside the symbols identify the data points used to determine the lines shown. Ea values for the selected sections of data are given near the data.

	DISCUSSION

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The main results are: (a) the rate of activation of the g_H is highly temperature sensitive. The Q₁₀ for both the delay and _act was about the same, ranging from 6 to 9 in various cells. (b) The rate of deactivation also has a high Q₁₀ of 6-8, practically identical to that for activation. (c) There were no obvious break points in the temperature dependence of the three kinetic parameters studied. (d) The Q₁₀ of the H⁺ conductance is high, >2 in whole-cell measurements, and 2.8 in inside-out patches >20°C and 5.3 at <20°C. (e) The Arrhenius plot of I_H is nonlinear, being steeper at low temperature. (f) The temperature dependence of both H⁺ currents and H⁺ channel gating kinetics is quite similar in different mammalian cells.

H⁺ Channel Gating Is Steeply Temperature Dependent

Table I summarizes the temperature dependence of three kinetic parameters reflecting channel opening (_act and delay) and closing (_tail). For all cells studied, the mean Q₁₀ values were similar, both from one cell type to another, and, more remarkably, for all three kinetic parameters. For other ion channels, opening and closing (_act and _tail) do not as a rule have the same Q₁₀ (Table II), and in most cases the Q₁₀'s of inactivation (or block) and recovery appear to be radically different. The Q₁₀ values in Table IA for H⁺ currents range from 6 to 9, which inspection of Table II shows is at the upper end of reported values for gating of various ion channels. Most channel gating processes have Q₁₀ values near 3, generally taken to indicate significant conformational rearrangement of the channel molecule. A high Q₁₀ for K⁺ channel inactivation was ascribed to interaction between two peptide moieties, at least one of which evidently has a low probability of adopting the correct conformation to permit binding (Murrell- Lagnado and Aldrich, 1993). Alamethicin pore formation (listed as _act in Table II) has a Q₁₀ of 9 at low temperatures. This result is intriguing because alamethicin is believed to form channels in which 6-10 molecules assemble in the membrane in a barrel-stave arrangement (Boheim and Kolb, 1978), reminiscent of one of the hypothetical physical depictions of our gating model, in which several channel protomers assemble in the membrane to form a functional channel (Cherny et al., 1995). Another intriguing parallel is the high Q₁₀ of a slow gating process in a Cl channel that, like the H⁺ channel, is gated by its permeant ion, interpreted as reflecting channel subunit interaction (Pusch et al., 1997). In general, the high Q₁₀ for H⁺ channel gating is compatible with substantial conformational changes in the channel.

We proposed a model of H⁺ channel gating in which opening requires deprotonation of an externally accessible site(s), followed by a conformational change that shifts the accessibility of the protonation site(s) to the internal side, and finally stabilization of the open configuration by protonation from the inside (Cherny et al., 1995). This model illustrated how gating could be regulated by voltage and the pH gradient, pH (pH_o  pH_i). Although the model is hypothetical, it is difficult to conceive of a mechanism that does not incorporate the general ideas of regulatory protonation sites whose accessibility switches from one side of the membrane to the other. The voltage dependence of gating in our model could arise either from voltage-dependent proton binding to sites inside "proton wells" analogous to those in H⁺ ATPases (Mitchell and Moyle, 1974) or from a voltage-dependent conformational change, as is more typical of voltage-gated ion channels. The protonation/deprotonation reactions at the internal site are rate determining, with the conformational change being rapid and in quasi-equilibrium. Because deuterium slowed activation threefold with only minor effects on deactivation, we suggested that a voltage-dependent closing step might precede the internal deprotonation step (DeCoursey and Cherny, 1997).

In this context, the similarity of Q₁₀ values for _act, delay, and _tail was surprising. Although we cannot rule out the possibility that several processes could have the same high E_a, the simplest explanation is that the same energy barrier is rate determining for all three parameters. The energy wells on either side of the barrier must be relatively symmetrical. The E_a for this process is evidently 30-38 kcal/mol, which is substantially larger than any of the proton-related processes listed in Table III. Specifically, the E_a is much larger than 7 kcal/mol for ionization of the imidazole group of histidine (Reeves, 1977), a candidate for the external regulatory protonation site, proposed on the basis of deuterium isotope effects on gating (DeCoursey and Cherny, 1997). In a linear gating scheme, the delay presumably reflects early events in the opening process, _act reflects the entire gating sequence, and _tail must reflect the final transition between the open state and a neighboring closed state. It appears paradoxical that the same process could determine the relatively rapid tail current decay and the slower activation. A possible explanation for the similarity of E_a for all three kinetic parameters is that each of several channel subunits must undergo an identical or similar complex first-order conformational change during opening, but a reverse transition in only one subunit is sufficient to close the channel. This is essentially the Hodgkin-Huxley (1952) model for Na⁺ and K⁺ channel gating. This hypothesis would also explain the surprising similarity of E_a for the delay, _act, and _tail. In a multiple independent subunit channel, the ratio of delay to _act is fixed (R.W. Aldrich and F.T. Horrigan, personal communication).

The temperature dependence of gating was nearly linear in Arrhenius plots. All three of the kinetic parameters measured had roughly linear temperature dependence over the range 6-42°C. For a constant Arrhenius activation energy, E_a, the temperature dependence in simple semi-logarithmic plots (e.g., Figs. 4 A, 5, and 7 A) is slightly nonlinear, becoming steeper at low temperatures. Our data do not resolve the fine distinction between linearity and the predicted Arrhenius curvature, and appeared linear on both types of plots (compare Fig. 4, A and B). Many studies of other ion channels describe break points in conductance (Lass and Fischbach, 1976; Chiu et al., 1979; Hagiwara and Yoshii, 1980; Quartararo and Barry, 1988) or gating kinetics (Schwarz, 1979; Chiu et al., 1979; Kirsch and Sykes, 1987), which occur at 4-20°C. Others report no clear break points, although the Q₁₀ increases at lower temperatures (Boheim and Kolb, 1978; Kimura and Meves, 1979; Kukita, 1982; Beam and Donaldson, 1983; Pahapill and Schlichter, 1990). Breakpoints in Arrhenius plots are generally taken to indicate phase transitions of membrane lipids. Krasne et al. (1971) reported that "freezing" artificial bilayer membranes abolished the conductance mediated by the carriers nonactin and valinomycin, but had little effect on the conductance of the gramicidin channel. Here we saw no consistent or reliable evidence of a break point in the temperature dependence of any kinetic parameter. The absence of a break point could mean that H⁺ channel gating is relatively insensitive to the fluidity of the surrounding membrane (see Beam and Donaldson, 1983). A more likely explanation is that the membranes of the cells studied do not exhibit sharp phase transitions in the temperature range studied. The linearity of the Arrhenius plots suggests that the same process is rate determining over the entire temperature range.

Arrhenius plots of I_H are nonlinear. In contrast to the kinetic parameters, I_H usually changed more steeply at low temperatures, and tended to saturate at high temperature (>30°C). In cells in which I_H saturated, the currents often decayed during the pulse (not shown). H⁺ channels do not inactivate, and this decay is almost certainly the result of increased pH_i due to the large efflux of H⁺ during current flow. Large I_H decreases the driving force by increasing pH_i, as has been demonstrated by changes in V_rev (DeCoursey, 1991; DeCoursey and Cherny, 1994b), or deduced from pH_i measured by microelectrodes (Thomas and Meech, 1982; Meech and Thomas, 1987) or fluorometric dyes (Kapus et al., 1993). During even moderate depolarizations at high temperature, there was sometimes evidence of depletion. Our interpretation is that high temperature exacerbates the depletion of protonated buffer from the cell (and consequent increase in pH_i), in spite of the high (100 mM) concentration of buffer in all solutions. Thus, the apparent saturation of I_H at high temperature is not ascribable to events occurring near the channel. Higher Q₁₀ values were obtained for I_H measured in inside-out patches than in whole-cell experiments, whereas Q₁₀'s for gating kinetics were similar. The most likely explanation is that depletion of protonated buffer due to H⁺ efflux is less problematic in excised patches because there are much smaller diffusion barriers between either side of the membrane and an effectively infinite volume of buffered solution.

I_H Has Abnormally Strong Temperature Dependence

The average Q₁₀ for I_H in whole-cell measurements was 2.1-3.1 in various mammalian cells, and was higher in patches (Table IA). For reasons discussed above, we consider Q₁₀ values from inside-out patches to be more reliable for I_H (but less reliable for the kinetic parameters). The Q₁₀ in patches averaged ~2.8 at >20°C and increased to 5.3 at <20°C (Table IA). As discussed in the context of Fig. 6, overestimation of the leak current would tend to decrease the apparent Q₁₀. Temperature effects on the pK_a of buffers will affect these values by changing both the absolute pH and pH. The decrease in pH_i at higher temperature will tend to increase I_H, presumably by ~1.7-fold/U (DeCoursey and Cherny, 1995, 1996a). Most of the measurements were made with Bes externally and Mes internally, for which pH will decrease at higher temperatures, essentially offsetting the increase in RT/F. Correcting for the decrease in pH_i at higher temperature lowers the Q₁₀ values to 2.0-2.9 in whole-cell and 2.6-5.0 in excised patch measurements. These values greatly exceed practically all values reported for ion permeation through other channels (Table II), and thus require some explanation.

An increase in I_H with temperature could reflect increased single-channel conductance or open probability, P_open. If the opening and closing rates had different temperature dependence, P_open would vary with temperature. Three observations indicate that P_open does not change with temperature. First, the g_H-V relationship was not convincingly shifted. Second, the kinetic parameters of gating (delay, _act, and _tail) and the I_H waveform in general all appeared to simply scale uniformly with temperature. The temperature dependence of neither _act nor _tail was detectably voltage dependent. Finally, the H⁺ current variance at 20°C increases sharply with depolarization, and then plateaus or decreases with further depolarization (V.V. Cherny and T.E. DeCoursey, unpublished observations), suggesting that for large depolarizations P_open > 0.8 at 20°C, which would severely limit any possible increase at higher temperatures by this mechanism. None of the arguments is conclusive, and none rules out mechanisms in which the number of functional channels changes with temperature. Insertion of channels at high temperature by vesicle fusion can be ruled out because lowering the temperature rapidly reverses the effects of temperature on I_H. Finally, we cannot eliminate the possibility that a rapid gating process (i.e., "flicker") that is perhaps not in the normal opening pathway might alter the effective unitary current with some arbitrary temperature dependence. In the discussion that follows, we assume that the temperature dependence of I_H reflects changes in unitary conductance.

The H⁺ current is not limited by bulk diffusion. Limitations to channel permeation can occur at several stages, as delineated for gramicidin by Andersen (1983). The rate-limiting step could occur during diffusion to the mouth of the channel, entry into the channel, permeation through the pore, the exit step, or diffusion away on the distal side. For H⁺ channels, additional possible rate-limiting steps include buffer diffusion, protonation/deprotonation reactions, and hydrolysis. Because varying the external or internal buffer concentrations from 1 to 100 mM changed I_H less than twofold (DeCoursey and Cherny, 1996b), neither diffusion nor protonation/deprotonation of buffer limits I_H. The present results appear to rule out the possibility that diffusion of free H⁺ is rate determining. This conclusion is consistent with the larger deuterium isotope effect on H⁺ channel currents than on conduction in bulk solution (DeCoursey and Cherny, 1997).

Temperature dependence of H⁺ conduction in water. The electrical conductivity (ionic mobility) of H⁺ is anomalously higher than that of any other cation, by a factor of ~5 (Eigen and DeMaeyer, 1958; Robinson and Stokes, 1959). This observation is explained by the unique conduction mechanism for H⁺. The proton in aqueous solution exists mainly (96-99% of the time) in association with a particular water molecule as a hydronium ion, H₃O⁺ (Conway et al., 1956). Any of the three protons may leave this molecule to conduct current. Protons thus hop from one water molecule to the next (de Grotthuss, 1806) by a mechanism referred to as Grotthuss, water wire, or prototropic transfer (Lengyel and Conway, 1983). The activation energy for H⁺ conduction is lower than for other ions, and decreases rapidly with increasing temperature. The Q₁₀ for H⁺ conductivity, ⁰, decreases from 1.2 between 5 and 15°C to 1.11 between 35 and 45°C (Landolt-Börnstein, 1960). The conductivity of H⁺ traditionally has been separated into ordinary hydrodynamic conduction (that expected if H₃O⁺ diffused as an invariant molecular species like other ions) and "excess" prototropic conductivity of H⁺; e.g., _H⁺  _Na⁺ (Hückel, 1928). Hydrogen bonds between molecules facilitate H⁺ conduction by the Grotthuss mechanism. The excess prototropic conduction decreases strongly with increased temperature: (_{HCl  KCl})/_KCl is 2.26 at 273°K and 1.07 at 373°K (Lengyel and Conway, 1983), because the extent of hydrogen bonding in water is decreased by thermal motion (Ewell and Eyring, 1937; Morgan and Warren, 1938). The validity of the common practice of subtracting hydrodynamic from total to obtain the "excess" H⁺ conductance has been questioned recently, on the basis that little conventional hydrodynamic conductance can occur due to the strength of first-shell hydrogen bonds, which trap the H₃O⁺ ion in a densely hydrogen-bonded network of water molecules (Agmon, 1996). In this view, all H⁺ conduction occurs by prototropic transfer. H⁺ permeation through channels most likely occurs by this mechanism.

Permeation is not equivalent to diffusion in bulk solution. The conductance of most cation channels is weakly temperature sensitive (Table II). A higher E_a has been reported for a lymphocyte K⁺ channel, 8.2 kcal/mol or twice that of free diffusion (Lee and Deutsch, 1990), and even higher values for inward rectifier K⁺ channels, although these higher values are from macroscopic measurements that may reflect factors other than changes in single-channel current amplitude. The Q₁₀ of the conductivity of physiological monovalent cations (Na⁺ and K⁺) in aqueous solutions is ~1.2-1.3 between 5 and 35°C, while that of H⁺ is only 1.14-1.20 (Robinson and Stokes, 1959). The similarity of the Q₁₀ for aqueous diffusion of ions and open channel conductance suggests that ions permeate channels in an environment approximating aqueous diffusion (e.g., Horn, 1984; Stein, 1986). This implies that there are not large energy barriers in the permeation pathway, and the ion should permeate at a rate roughly comparable with its diffusion in bulk solution. The conductance of various channels is somewhat less than calculated from the bulk diffusion coefficient, given the dimensions of the pore (Stein, 1986). Calculated using the Poisson-Nernst-Planck approach, the effective diffusion coefficient of ions inside channels during permeation is roughly an order of magnitude lower than in bulk solution (Chen et al., 1997). Part of the higher E_a for conduction through ion channels than bulk solution may reflect the energetic cost of partial dehydration of the ion; the effective hydration number of ions in solution decreases at higher temperatures, facilitating entry into the channel (Kuyucak and Chung, 1994). The mobility of H⁺ in gramicidin channels, which are single-file water-filled pores (Levitt et al., 1978; Finkelstein and Andersen, 1981), at high [HCl] is not much lower than in bulk HCl solution, suggesting that this water-filled pore offers little additional intrinsic resistance (Cukierman et al., 1997). H⁺ current through single gramicidin channels has a Q₁₀ of 1.33 (4.8 kcal/mol) at 1 M HCl (Akeson and Deamer, 1991), about twice that of the mobility of H⁺ in bulk solution. Akeson and Deamer (1991) speculated that misaligned hydrogen bonds between waters in the pore might increase the E_a for proton hopping. The recent proposal that the rate-determining step in H⁺ conduction in water is the breaking of second hydration shell hydrogen bonds of strength 2.5 kcal/mol (Agmon, 1995) leads to the idea that entry of H⁺ into a channel must require breaking a first shell hydrogen bond, which would be about twice as strong (Agmon, 1996), consistent with the conclusion that H⁺ permeation through gramicidin is entirely diffusion limited (Decker and Levitt, 1988). The Q₁₀ for H⁺ permeation reported here is substantially larger, especially considering the uniquely low Q₁₀ of H⁺ conduction in aqueous solution. Evidently, the rate-limiting step in H⁺ permeation through voltage-gated H⁺ channels is thermodynamically distinct from diffusion.

What is the rate-limiting step in permeation? Table III compares E_a for voltage-gated H⁺ channels with various processes involving protons. We focus on parameters related to hydrogen bonding an