The voltage-activated H+ selective conductance of rat alveolar epithelial cells was studied using whole-cell and excised-patch voltage-clamp techniques. The effects of substituting deuterium oxide, D2O, for water, H2O, on both the conductance and the pH dependence of gating were explored. D+ was able to permeate proton channels, but with a conductance only about 50% that of H+. The conductance in D2O was reduced more than could be accounted for by bulk solvent isotope effects (i.e., the lower mobility of D+ than H+), suggesting that D+ interacts specifically with the channel during permeation. Evidently the H+ or D+ current is not diffusion limited, and the H+ channel does not behave like a water-filled pore. This result indirectly strengthens the hypothesis that H+ (or D+) and not OH− is the ionic species carrying current. The voltage dependence of H+ channel gating characteristically is sensitive to pHo and pHi and was regulated by pDo and pDi in an analogous manner, shifting 40 mV/U change in the pD gradient. The time constant of H+ current activation was about three times slower (τact was larger) in D2O than in H2O. The size of the isotope effect is consistent with deuterium isotope effects for proton abstraction reactions, suggesting that H+ channel activation requires deprotonation of the channel. In contrast, deactivation (τtail) was slowed only by a factor ≤1.5 in D2O. The results are interpreted within the context of a model for the regulation of H+ channel gating by mutually exclusive protonation at internal and external sites (Cherny, V.V., V.S. Markin, and T.E. DeCoursey. 1995. J. Gen. Physiol. 105:861–896). Most of the kinetic effects of D2O can be explained if the pKa of the external regulatory site is ∼0.5 pH U higher in D2O.
Voltage-gated H+ channels conduct H+ current with extremely high selectivity and exhibit voltage-dependent gating that is strongly modulated by both extracellular and intracellular pH (pHo and pHi, respectively).1 Here we explore the effects of substituting heavy water (deuterium oxide, D2O), for water (protium oxide, H2O), on both the conductance and the pH dependence of channel gating. The isotope effect on conductance should provide insight into the mechanism by which permeation occurs. Isotope effects on the regulation by pH (or pD) of the voltage dependence and kinetics of gating provide clues to the possible protonation/deprotonation reactions that have been proposed to play a role in channel gating (Byerly et al., 1984; Cherny et al., 1995).
Several chemical properties of D2O and H2O are compared in Table I. From the perspective of this study, the main differences between D2O and H2O are: (a) the viscosity of D2O is 25% greater than H2O, (b) the conductivity of H+ in H2O is 1.4–1.5 times that of D+ in D2O, (c) H+ has a much greater tendency than D+ to tunnel, (d) D+ weighs twice as much as H+, and (e) D+ is bound more tightly in D3O+ and in many other compounds than is H+. Three main types of deuterium isotope effects are recognized: general solvent effects and primary and secondary kinetic effects. General solvent isotope effects reflect the different properties of D2O and H2O as solvents, such as viscosity or dielectric constant. As seen in Table I, these differences are rather moderate, and their effects are accordingly usually moderate as well. Kinetic isotope effects reflect involvement of protons or deuterons in chemical reactions. Primary kinetic isotope effects occur when H+ directly participates in a rate-determining step in the reaction, for example a protonation/deprotonation or H+ transfer reaction. For example, ionization of a number of bases is typically three to seven times slower in D2O (Bell, 1973). Secondary isotope effects reflect D+ for H+ substitution at some site distinct from the primary reaction center. Secondary kinetic isotope effects tend to be small, 1.02–1.40 (Kirsch, 1977).
The conductivity of H+ is about five times higher than that of other cations with ionic radii like that of H3O+; the limiting equivalent conductivity (λ0) at 25°C in water is 350 S cm2/equiv. for H+ but 73.5 S cm2/equiv. for NH4+ (Robinson and Stokes, 1965). This anomalously high conductivity for H+ has been ascribed to conduction by a mechanism in which H+ jumps from H3O+ to a neighboring water molecule (Danneel, 1905; Hückel, 1928; Bernal and Fowler, 1933; Conway et al., 1956). H+ hopping can occur faster than ordinary hydrodynamic diffusion (i.e., bodily movement of an individual H3O+ molecule analogous to the diffusion of ordinary ions). After one H+ conduction event, a structural reorientation of the hydrogen-bonded water lattice is necessary before another proton can be conducted (Danneel, 1905; Bernal and Fowler, 1933; Conway et al., 1956). Proton conduction through channels is believed to occur by an analogous two-step “hop-turn” process through a hydrogen-bonded chain or “proton wire” spanning the membrane (Nagle and Morowitz, 1978; Nagle and Tristram-Nagle, 1983).
The mobility (measured as conductivity) of H+ in H2O is 1.41 times that of D+ in D2O (Table I); nevertheless, the mobility of D+ is still 4 times that of K+ in D2O (Lewis and Doody, 1933). Thus, D+ also exhibits abnormally large conductivity, even though tunnel transfer of D+ is 20 times less likely than for H+ and one might have expected simple hydrodynamic diffusion of D3O+ to play a larger role for D+, which would accordingly have a conductivity similar to that of other cations (Bernal and Fowler, 1933). Evidently the reorientation of hydrogen-bonded water molecules (the turning step of a hop-turn mechanism) is rate limiting for both H+ and D+ conduction. The nature of this rate-determining step has been proposed to be the reorientation of hydrogen-bonded water molecules in the field of the H3O+ ion (Conway et al., 1956), “structural diffusion” or formation and decomposition of hydrogen bonds at the edge of the H9O4+ complex (i.e., the hydronium ion with its first hydration shell) (Eigen and DeMaeyer, 1958), or more recently, the breaking of an ordinary second-shell hydrogen bond converting H9O4+ to H5O2+ (Agmon, 1995, 1996). Some such reorganization of hydrogen bonds may also be the rate limiting step in proton translocation across water-filled ion channels such as gramicidin (Pomès and Roux, 1996).
A characteristic feature of voltage-gated H+ currents is their sensitivity to both pHo and pHi. Increasing pHo and decreasing pHi shift the voltage-activation curve to more negative potentials in every cell in which these parameters have been studied (reviewed by DeCoursey and Cherny, 1994). This effect of pH is reminiscent of its effects on many other ion channels, which may reflect the neutralization of negative surface charges (see Hille, 1992). However, the magnitude of the pH-induced voltage shifts for H+ currents has led to the suggestion that protonation of specific sites on or near the channel allosterically modulate gating (Byerly et al., 1984). In alveolar epithelial cells (Cherny et al., 1995), as well as in other cells (DeCoursey and Cherny, 1996a; Cherny et al., 1997), the shift produced by internal and external protons (H+i and H+o) is quite similar, 40 mV/U change in ΔpH, within a large pH range encompassing physiological values. Thus the position of the voltage activation curve can be predicted from the pH gradient, ΔpH, rather than by pHo and pHi independently. This behavior was explained by a model (Cherny et al., 1995) in which there exist similar protonation sites accessible from either the internal or external solution, but not both simultaneously. Protonation from the outside stabilizes the closed channel, whereas protonation from the inside stabilizes the open channel. Here we show that H+ channels are regulated in a similar manner by D+, but that D+ binds more tightly to the modulatory sites on the channel molecule.
Materials And Methods
Alveolar Epithelial Cells
Type II alveolar epithelial cells were isolated from adult male Sprague-Dawley rats under sodium pentobarbital anesthesia using enzyme digestion, lectin agglutination, and differential adherence, as described in detail elsewhere (DeCoursey et al., 1988; DeCoursey, 1990). Briefly, the lungs were lavaged to remove macrophages, elastase and trypsin were instilled, and then the tissue was minced and forced through fine mesh. 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 become more like type I cells. No obvious changes in the properties of H+ currents have been observed. H+ currents were studied in approximately spherical cells up to several weeks after isolation.
Most solutions (both external and internal) contained 1 mM EGTA, 2 mM MgCl2, 100 mM buffer, and TMAMeSO3 added to bring the osmolarity to ∼300 mosM, and titrated to the desired pH with tetramethylammonium hydroxide or methanesulfonic acid (solutions using BisTris as a buffer). The pH 8, 9, and 10 solutions contained 3 mM CaCl2 instead of MgCl2. A stock solution of TMAMeSO3 was made by neutralizing tetramethylammonium hydroxide with methanesulfonic acid. Buffers (Sigma Chemical Co., St. Louis, MO), which were used near their pK in the following solutions, were: pH 5.5, pD 6.0 Mes; pH 6.5, pD 7.0 Bis-Tris (bis[2-hydroxyethyl]imino-tris[hydroxymethyl]methane); pH 7.0, pD 7.0 BES (N,N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid); pH 7.5, pD 8.0 HEPES; pH 8.0 Tricine (N-tris[hydroxymethyl] methylglycine); pH 9.0, pD 9.0 CHES (2-[N-cyclohexylamino] ethanesulfonic acid); pH 10, pD 10 CAPS (3-[cyclohexylamino]- 1-propanesulfonic acid). The pH (or pD) of all solutions was checked frequently.
A series of solutions containing NH4+ was made to impose a defined pH gradient across the cell membrane, as described by Grinstein et al. (1994). The principle is that if neutral NH3 molecules permeate the membrane rapidly enough to approach identical concentrations on both sides of the membrane, then:
because the bath solution is heavily buffered (100 mM buffer) and diffuses freely but the pipette solution (for these measurements) is weakly buffered and diffusion is slowed by the pipette tip. The shift of pHi occurs because [H+]i = pKa − log [NH4+]i/ [NH3]i. The extracellular solutions were made with 100 mM HEPES, 2 mM MgCl2, 1 mM EGTA, and various concentrations of (NH4)2SO4, at pH 7.5. TMAMeSO3 was added to bring the osmolarity to ∼300 mosM. The pipette solution, which was also used externally, included 25 mM (NH4)2SO4, 5 mM BES, 2 mM MgCl2, 1 mM EGTA, and TMAMeSO3, brought to pH 7.0 with tetramethylammonium hydroxide.
We assume that when NH4+ diffuses from the pipette into the cell, if D2O is present in the bath (and hence inside the cell) there will be rapid exchange of D+ for H+ in NH4+, and that therefore efflux of ND3 will occur, leaving D+ rather than H+ behind inside the cell. Deuterons in deutero-ammonia, ND3, exchange rapidly with protons (Cross and Leighton, 1938).
The osmolarity of solutions was measured with a Wescor 5500 Vapor Pressure Osmometer (Wescor, Logan, UT). Deuterium oxide (99.8% or 99.9%) was purchased from Sigma Chemical Co. A liquid junction potential of ∼2 mV was measured between solutions identical except that D2O replaced H2O. If water did not permeate the cell membrane, correction for this junction potential would make the transmembrane potential 2 mV more negative. However, as described in Fig. 1, we feel that water permeates the cell membrane, and thus there would be offsetting junction potentials at the pipette tip and bath electrode even in whole cell configuration. Therefore no junction potential correction has been applied.
The reading taken from a glass pH electrode, pHnom, deviates from the true pD of D2O solutions by 0.40 U, such that pD = pHnom + 0.40 (Glasoe and Long, 1960). Another estimate of this difference is 0.45 ± 0.03 (Dean, 1985), and even more disparate values can be found in early studies. Given the uncertainty about the precise value, we tested our pH meter (Radiometer Ion83 Ion meter; Radiometer, Copenhagen, Denmark) following the approach taken by Glasoe and Long (1960). Our pH meter read 0.402 ± 0.006 (mean ± SD, n = 3) higher when 0.01 M HCl was added to H2O than when added to D2O. We therefore corrected the pD in D2O solutions by adding 0.40 to the nominal reading of our pH meter.
Estimation of the pKa of the Buffers in H2O and in D2O
Most simple carboxylic and ammonium acids with pKa between 4 and 10 have a pKa 0.5–0.6 U higher in D2O than in H2O (Schowen, 1977). We titrated the buffers used in this study at room temperature (20–23°C). 10 mmol of buffer was added to 20 ml of H2O or D2O and titrated with 10 N NaOH, or 10 N HCl in the case of Bis-Tris. The resulting contamination of D2O by the H+ from the base or acid titrating solutions is <3%. We corrected for this error in two ways. First, we increased the apparent change in pKa, assuming a linear mole-fraction dependence (cf. Glasoe and Long, 1960), which increased the pKa in D2O by ≤0.02 U. We also carried out some titrations using deuterated acids and bases (DCl and NaOD, both from Aldrich Chemical Co, Milwaukee, WI). The results by these two methods were similar. The averages of two to three separate determinations for each buffer are given in Table II.
Conventional whole-cell, cell-attached patch, or excised inside-out patch configurations were used. Experiments were done at 20°C, with the bath temperature controlled by Peltier devices and monitored continuously by a thinfilm platinum RTD (resistance temperature detector) element (Omega Engineering, Stamford, CT) immersed in the bath. Micropipettes were pulled in several stages using a Flaming Brown automatic pipette puller (Sutter Instruments, 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 3–10 MΩ. 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 Corporation, Sunnyvale, CA). Data acquisition and analysis programs were written in BASIC-23 or FORTRAN. Seals were formed with Ringer's solution (in mM: 160 NaCl, 4.5 KCl, 2 CaCl2, 1 MgCl2, 5 HEPES, pH 7.4) 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.
For “typical” families of H+ currents, pulses were applied in 20-mV increments with an interval of 30–40 s or more, depending on test pulse duration and the behavior of each particular cell. Although 30 s is not long enough for complete recovery from the depletion of intracellular protonated buffer, it represents a compromise aimed at allowing multiple measurements to be made in each cell reasonably close together in time. For some measurements in which only small currents were elicited, such as pulses in 5-mV increments near Vthreshold, a smaller interval between pulses was used, because negligible depletion was expected. We tried to bracket measurements in different solutions whenever possible.
The time constant of H+ current activation, τact, was obtained by fitting the current record by eye with a single exponential after a brief 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 tdelay is the delay. The H+ current amplitude is (I0 − IΘ). No other time-dependent conductances were observed consistently under the ionic conditions employed. Tail current time constants, τtail, were fitted either to a single decaying exponential:
where I0 is the amplitude of the decaying part of the tail current, or to the sum of two exponentials:
where An are amplitudes and τn are time constants.
We refer to the pL in the format pLo//pLi. In the inside-out patch configuration the solution in the pipette sets pLo, which is defined as the pL of the solution bathing the original extracellular surface of the membrane, and the bath solution is considered pLi. 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.
As discussed in detail in Strategic Considerations and in Fig. 1, when the bath solvent differs from that in the pipette, the effective pHi (or pDi) will differ from the nominal value of the pipette solution by ∼0.5 U. Therefore, when bath and pipette solvents differ, we provide values for the presumed effective internal H+ or D+ concentration, e.g., pHi,eff 6.5 indicates a pD 7.0 pipette solution with any H2O solution in the bath. The majority of experiments were done with D2O rather than with H2O pipette solutions because we wanted the measurements in D2O to be contaminated as little as possible by H2O.
The nature of the problem under investigation introduces several complications, which require explanation, as well as a perhaps less-than-obvious approach. Ideally we would like to compare the behavior of the proton conductance in the same cell under identical conditions while varying only the solvent (D2O or H2O) on one side of the membrane and keeping pLo and pLi constant (pLx refers to either pHx or pDx). However, the high membrane permeability of water means that only symmetrical solvent studies can be contemplated. Less obviously, due to the increased pKa of buffer in D2O (Table II), it is impossible to compare directly in the same cell identical pHo and pDo by simply changing the external solvent, without at the same time changing pLi. However, it is desirable to make comparisons in the same cell, because H+ currents vary substantially from cell to cell. We therefore adopted two strategies. First, we compare currents measured with the same pH or pD gradient (e.g., pHo 6.5//pHi 6.5 and pDo 7.0// pDi 7.0), because the gradient, ΔpH, appears to be a fundamental determinant of H+ channel gating (Cherny et al., 1995). This approach has the drawback of comparing the effects of different absolute concentrations of protons and deuterons, and there is some indication that H+ channel gating kinetics depend on the absolute pHi, rather than ΔpH alone (DeCoursey and Cherny, 1995). The second approach (see materials and methods) overcomes this shortcoming by controlling pHi by applying a known NH4+ gradient (Roos and Boron, 1981), as illustrated by Grinstein et al. (1994). Varying the NH4+ gradient allows resetting pHi (or pDi) in a cell under whole-cell voltage-clamp, and ideally, comparison of currents at the same pH and pD.
Only symmetrical solvent is possible.
In these experiments we varied the solvent in the pipette and bath solutions. Because water has a high membrane permeability, it seemed likely that the solvent in the bath solution would enter the cell much faster than solvent would diffuse from the pipette, and thus the solvent in the bath would also be present in the cell, regardless of the pipette solution. This expectation was tested theoretically and experimentally.
• How fast does water enter the cell?
A critical question in the interpretation of the data is whether solvent in the bath diffuses across the cell membrane fast enough to dominate the intracellular solution in spite of the presence of the pipette tip which is a continuous source of solvent from the pipette solution. The water permeability, Posm, of planar lipid bilayers or liposomes ranges from 10−4 cm/s to 10−2 cm/s; Posm in various epithelial cell membranes similarly ranges from 10−4 cm/s to >10−2 cm/s (Tripathi and Boulpaep, 1989). Because both HgCl2-sensitive and HgCl2-insensitive water channels occur in lung tissue (Folkesson et al., 1994; Hasegawa et al., 1994), it is likely that Posm is relatively high in alveolar epithelial cells, at least in situ. Osmotic water permeability (Pf) is 1.7 ± 10−2 cm/s and diffusional water permeability, Pd, is 1.3 ± 10−5 cm/s across the alveoli of intact mouse lung (Carter et al., 1996). However, Pd was probably grossly underestimated because of unstirred layer effects (Finkelstein, 1984; Carter et al., 1996). We calculated the steady-state distribution of normal or heavy water when one species was in the pipette solution and the other in the bath solution. The compartmental diffusion model used has been described in detail previously (DeCoursey, 1995), and simplifies the calculation by placing the pipette tip at the center of a spherical cell. The diffusion coefficient of H2O was taken as 2.1 × 10−5 cm2/s (Robinson and Stokes, 1965), the pipette tip was assumed to have a diameter of 1.0 μm, the cell diameter was 20 μm, and we assume that D2O and H2O have similar membrane permeabilities (Perkins and Cafiso, 1986; Deamer, 1987; Gutknecht, 1987). A range of Posm was assumed. For Posm > 10−3 cm/s the membrane presented essentially no barrier to diffusion, and the solvent in the bath was the main solvent inside the cell. Nevertheless, because the pipette is a constant source, there is always a finite concentration of the pipette solvent. For the pipette tip at the center of a 20 μm diameter cell, the limiting submembrane concentration at infinite Posm is ∼2% due to that in the pipette. Lowering Posm to 10−4 cm/s caused the membrane to become a significant diffusion barrier, with the steady-state concentration of solvent near the inside of the membrane 24% due to the pipette and 76% due to the bath. The fraction of solvent near the membrane originating in the pipette would be larger in a smaller cell but would be smaller if the pipette tip diameter were smaller. In conclusion, the pipette solvent is present in the cell at significant levels only for a quite conservative estimate of Posm, and in all likelihood the solvent in the bath permeates the membrane rapidly enough that most of the solvent near the membrane originated in the bath. We therefore assume that the membrane is exposed to nearly symmetrical solvent, with a finite but small contribution from the pipette.
• What is the pL (pH or pD) inside the cell?
The actual pLi can be deduced from knowledge of pLo and the reversal potential, Vrev. In the experiment illustrated in Fig. 1, the pipette contained pD 7.0 solution, and the tail current reversal potential, Vrev, was measured in several different bath solutions. Vrev was near 0 mV when the bath contained pD 7.0 (Fig. 1,A) or pH 6.5 (Fig. 1,C), and was −27 mV at pHo 7.0 (Fig. 1,B). In eight cells, Vrev was 29.9 ± 4.5 mV (mean ± SD) more negative at pHo 7.0 than at pDo 7.0, both with pDi 7.0. Reversal near 0 mV is expected for symmetrical pD 7.0//7.0. Why was Vrev near 0 mV at pHo 6.5 but not at pHo 7.0, under nominally symmetrical bi-ionic conditions? The explanation arises from the fact that many molecules bind D+ more tightly than H+. Most simple carboxylic and ammonium acids with pKa between 4 and 10, including buffers, have a pKa 0.5–0.6 U higher in D2O than in H2O (Schowen, 1977). We confirmed this generalization by titrating the buffers used in this study in both H2O and D2O and found pKa shifts ranging 0.60– 0.69 U (Table II). Fig. 1,D illustrates diagrammatically the effect of this pKa difference on a cell studied in the whole-cell configuration. The cell nominally contains the pipette solution with its buffer titrated to some pH or pD, in this example pD 7.0. If the solvent in the bath differs from that in the pipette, the bath solvent will replace the pipette solvent inside the cell, as discussed above. Because H+ has a lower affinity for buffer than does D+, fewer H+ will be bound to buffer than were D+, and hence the actual pHi will be lower by ∼0.5 U than was the pD of the pipette solution. This is true regardless of the actual value of pHo, because it results from the solvent dependence of the pKa of the buffer. The chart in Fig. 1 summarizes the experiment illustrated. Given the bath and pipette solutions, the observed Vrev agrees well with EH calculated with the assumptions that (a) the solvent in the bath completely replaces that in the cell, and (b) the effective pHi will be ∼0.5 U lower than pD in the pipette when H2O replaces D2O in the bath. By similar logic, when H2O is in the pipette solution and D2O is in the bath, the actual pDi will be ∼0.5 U higher than pHi with H2O in the bath.
• Vrev measurements are consistent with high water permeability and the 0.5 U pKa correction for intracellular buffer in D2O
We proposed above that the bath solvent will “fill” the cell regardless of the pipette solvent and that when the bath solvent differs from that in the pipette, pLi will change by ∼0.5 U from its nominal value. To a first approximation these assumptions seem reasonable, but two possible sources of error should be considered. First, some finite fraction of solvent in the cell is derived from the pipette. We could not determine from our data the extent of this “contamination.” Second, we assume that the buffer pKa increases exactly 0.5 U when D2O replaces H2O, although the true change may be slightly higher and may differ for different buffers. Our titration of several buffers used (Table II) revealed an average pKa shift of 0.67 U in D2O. To test the adequacy of our approximation of a 0.5 U shift, we compared the value for Vrev measured in the same cell in D2O and in H2O at 0.5 U lower pLo. The difference in Vrev averaged 2.9 ± 0.7 mV (mean ± SEM, n = 21) for pDo 6.0–pHo 5.5, pDo 7.0–pHo 6.5, and pDo 8.0–pHo 7.5. We could not detect any significant difference between buffers in this respect. By this measure the actual pHi may be ∼0.05 U more acidic than our assumed value, i.e., pHi may be 0.55 U lower than pDi. However, considering that the slope of the Vrev vs. ΔpH relationship in water was 52.4 mV (Cherny et al., 1995) compared with 58.2 mV for EH, possibly indicating a ∼10% attenuation of the ΔpH applied across the membrane, one might suggest that the change in buffer pKa should also be attenuated by 10% for internal consistency.
A complementary comparison can be made between Vrev measured in the same bath solution, but with H2O or D2O in the pipette solution. At pDo 7, Vrev averaged +4.5 ± 1.2 mV (mean ± SEM, n = 4) with pHi 6.5 and +4.3 ± 0.8 mV (n = 12) with pDi 7. At pHo 6.5, Vrev averaged +2.0 ± 1.6 mV (n = 4) with pHi 6.5, and +0.5 ± 1.1 mV (n = 10) with pDi 7. Thus, no systematic difference was observed in Vrev with D2O or H2O in the pipette. Together these data support the validity of the assumptions used to interpret these experiments.
Reversal Potential of D+ Currents
Values of Vrev obtained from tail current measurements, such as those illustrated in Fig. 1, A–C, in bilateral D2O are plotted as a function of the pD gradient in Fig. 2. In most experiments, Vrev was slightly positive to the calculated Nernst potential for D+, ED (dark line), reminiscent of the small positive deviations of Vrev from EH reported in most studies of H+ currents. Most of the data points for each pDi parallel ED, clearly establishing the selectivity of this conductance for D+. The largest deviation occurred at pDo 10//pDi 8. Parallel experiments in H2O solutions (not shown) produced a similar but more exaggerated result—Vrev followed EH closely up to pHo 8, with a smaller shift at pHo 9, and no further shift at pHo 10. The simplest interpretation of this result is that at high pHo there is a loss of control over pHi.
A more traditional but less attractive interpretation of the deviations of Vrev from ED is that the selectivity of the conductance for D+ is not absolute, and that at high pL the permeability to some other ion (e.g., TMA+) is increased. However, the observed deviations are not consistent with a constant permeability of TMA+ relative to D+, because they were roughly the same at a given pD gradient, ΔpD, at various absolute pD. Thus, the ratio PTMA/PD calculated using the GHK voltage equation was 2 × 10−7, 2 × 10−8, and 5 × 10−9 at pD · 6, pD · 7, or pD · 8, respectively, all at ΔpD = 2.0. Barring a bizarrely concentration-dependent permeability ratio, it appears that the conductance is extremely selective for D+ (or H+), with a relative permeability >108 greater for D+ than for TMA+.
Behavior of the Proton Conductance in D2O
Effects of changes in pDo.
After complete replacement of water with heavy water, D+ currents behaved qualitatively like H+ currents in normal water. Typical families of currents are illustrated in Fig. 3, with pDi 6 and pDo 8, 7, or 6. At relatively negative potentials only a small time-independent leak current was observed. During depolarizing pulses a slowly activating outward current appeared. The current has a sigmoid time course, and activation was faster at more positive potentials. Decreasing pDo produced two distinct effects on the currents. The voltage at which the conductance was first activated, Vthreshold, became more positive by about 40 mV/U decrease in pDo, and the rate of current activation became slower. This shift in the position of the voltage-activation curve is more apparent in Fig. 4. The currents measured at the end of 8-s pulses are plotted (solid symbols), as well as the amplitude extrapolated from a single-exponential fit to the rising phase (open symbols). This latter value corrects for the fact that the currents did not always reach steady state by the end of the pulses, as well as correcting for any time-independent leak current. In this example, and in other experiments, the shift in the current-voltage relationship was very nearly 40 mV/U decrease in pDo. These effects are quite similar to those of changes in pHo in water (Cherny et al., 1995).
Another effect of changes in pDo evident in Fig. 3 is that the conductance was activated more slowly at lower pDo. The time course of activation of H+ or D+ currents was fitted by a single exponential after a delay (Eq. 2). In some cases the fit was good, as in the example shown in the inset to Fig. 5, but sometimes the time course was more complex, with fast and slow components. Deviations from an exponential time course seemed most pronounced at large positive voltages and when there was a large pD gradient. Activation time constants, τact, in the same cell at pDo 8, 7, and 6 are plotted in Fig. 5. At each pDo τact is clearly voltage dependent, decreasing with depolarization. Lowering pDo appears to shift the τact-V relationship to more positive potentials and upwards, slowing activation in addition to shifting the voltage dependence. Similar results were obtained in other cells. Although the magnitude of τact varied from cell to cell, the effects of changes in pDo in each cell were quite similar to those illustrated.
Effects of changes in pDi.
The effects of pDi on D+ currents were studied both in whole-cell experiments and in excised patches. Studying patches allows a direct comparison in the same membrane. Fig. 6 illustrates D+ currents in an inside-out patch at pDo 8.0 and pDi 6.0 (A) or pDi 7.0 (B). In this and in several other patches Vthreshold was shifted by about −40 mV/U decrease in pDi. Time-dependent outward current first appeared at −40 mV at pDi 6.0 and at 0 mV at pDi 7.0. The small amplitude of most patch currents in D2O limits the quantitative accuracy of any conclusions. However, the conductance approximately doubled when pHi was reduced 1 U, comparable with the 1.7-fold increase/U decrease in pHi reported previously in inside-out patches (DeCoursey and Cherny, 1995). It is also obvious that activation was much faster at lower pDi.
The effects of changes in pDi in whole-cell experiments were explored in individual cells by varying the NH4+ gradient across the cell membrane (materials and methods). Fig. 7 illustrates families of D+ currents in a cell at two NH4+ gradients. In each case pDo was 7.5, but pDi decreased as the NH4+ in the bath was lowered. With a 1//50 NH4+ gradient (A) Vrev was −66 mV, and with a 15//50 NH4+ gradient (B) Vrev was −27 mV. On the basis of this change in Vrev, pDi was ∼0.7 U lower in A than in B. At lower pDi the currents activated more rapidly and the conductance appeared to be increased. Qualitatively similar effects of changes in pHi were seen in H2O solutions at various NH4+ gradients in alveolar epithelium (not shown) and in macrophages (Grinstein et al., 1994).
Deuterium Isotope Effects on H+ (D+) Currents
Voltage-gated current amplitude.
The average ratios of the current measured in individual cells both in effectively symmetrical H2O and symmetrical D2O are plotted in Fig. 9. The “steady-state” current amplitudes were obtained by extrapolation of single exponential fits (Eq. 2). At all potentials the currents were substantially larger in H2O. The ratio decreased at more positive potentials, but two sources of error would tend to cause a voltage-independent effect to deviate in this direction. First, during large depolarizations there is depletion of protonated (or deuterated) buffer from the cell, which tends to reduce the currents in a current-dependent manner. Because the currents were larger in H2O, there would be more attenuation than in D2O. Second, to the extent that the position of the voltage-activation curve may be shifted slightly positive in D2O relative to H2O (e.g., see Figs. 10 and 11), a smaller fraction of the total conductance would be activated in D2O, and this would mainly affect smaller depolarizations to the steep part of the gH-V relationship. Thus, it is not clear whether this effect was voltage dependent. The average ratio at +80 and +100 mV was 1.92 at pD 8 compared with pH 7.5, 1.91 at pD 7 compared with pH 6.5, and 1.65 at pD 6 compared with pH 5.5. In summary, the current carried by H+ through proton channels is about twice as large as that carried by D+.
Comparison of the g H -voltage and g D-voltage relationships.
In symmetrical D2O the conductance-voltage relationship shifted about 40 mV/U change in ΔpD just as in H2O. However, the absolute voltage dependence might be different in the two solvents. To address this possibility we compared similar ΔpH and ΔpD in the same cell, varying the NH4+ gradient to regulate pLi. Fig. 10 illustrates a typical experiment. Measurements were made in D2O (filled symbols) and in water (open symbols) at 1// 50 NH4+ (▴), 3//50 mM NH4+ (♦), and 15//50 mM NH4+ (▪). At each NH4+ gradient, the gD-V relation was shifted 10–15 mV positive to the corresponding gH-V relation. Moreover, Vrev was consistently more positive in D2O at any given NH4+ gradient. Apparently NH4+ gradients were less effective at clamping pLi in D2O, perhaps reflecting the higher viscosity of D2O (Table I), or the higher pKa of NH4+ in D2O (Lewis and Schutz, 1934)—at any given pL there would be a smaller concentration of neutral ND3 than NH3 available to permeate the membrane. The cytoplasmic acidifying power of 3 mM NH4+ in D2O might be roughly equivalent to that of 1 mM NH4+ in H2O, as was observed in the experiment illustrated in Fig. 10, if the neutral form were present at equal concentration, because the NH4+ gradient changes pLi in a dynamic manner through a sustained flux of neutral NH3. Indeed, Grinstein et al. (1994) found that methylamine+, with a pKa 10.19 compared with 9.24 for NH4+ (Dean, 1985), acidified the cytoplasm more slowly given the same gradient than did NH4+. If one assumes that Vrev accurately reflects pLi then correcting for the difference in Vrev reduces the average shift in D2O (compared with H2O) to only ∼5 mV. Scaling the D2O data up to correct for the smaller limiting conductance further reduces the size of the shift. A residual shift of a few mV cannot be ruled out, but any such shift is not large, and it is possible that there is no shift.
Fig. 10 also shows that the conductance near threshold potentials changed e-fold in 4–5 mV at each NH4+ gradient. We could not detect any difference in this limiting slope in D2O and H2O. Measured at 10−2 to 10−3 of its maximal value, the conductance changed e - fold in 4.65 ± 0.16 mV (mean ± SEM, n = 22) in D2O and H2O combined; the lines drawn through the data in Fig. 10 illustrate this average slope. This slope corresponds with the translocation of 5.4 charges across the membrane during gating, which should be considered a lower bound for the actual gating charge movement.
Finally, examination of the limiting maximum conductance at large depolarizations (Fig. 10) reveals that over the range of pLi studied, the conductance was about twice as large in H2O as in D2O. This result is an important corroboration of the conclusion drawn from Figs. 8 and 9, because those comparisons were at ∼0.5 U different absolute pLi. The higher conductance in H2O than in D2O in Fig. 10 cannot be ascribed to different pLi and must reflect a fundamental difference in the rate at which D+ and H+ permeate the channel.
Relationship between Vthreshold and Vrev.
The potential at which the H+ conductance is first activated by depolarization, Vthreshold, is plotted in Fig. 11 as a function of Vrev in H2O (open symbols) and in D2O (filled symbols). Data obtained at pHo 6.5–10.0 and pDo 7–10 are included, as well as from experiments in which pLi was changed by varying the NH4+ gradient across the membrane. The data describe a remarkably linear relationship, with no suggestion of saturation at either extreme. The data for effectively symmetrical H2O and D2O fitted independently by linear regression yielded identical slopes (0.76 for H2O and 0.75 for D2O). Thomas (1988) observed a similarly linear relationship between EH and Vrev in snail neurons, over a range of pHi ∼7–8. This result shows clearly that the fundamental determinant of the position of the voltage-activation curve of the gH is the pH gradient across the membrane, as was concluded previously (Cherny et al., 1995).
The regression line in Fig. 11 for D2O is shifted 3.9 mV from that for H2O, indicating a more positive Vthreshold for a given Vrev. This small shift may be an artifact resulting from the greater difficulty in detecting small currents in D2O because the conductance is smaller and activation is slower. In any case, there was little or no solvent dependence of the relationship between Vrev and Vthreshold, suggesting the position of the voltage-activation curve of the proton conductance is fixed in a very similar manner by ΔpD as by ΔpH.
Deuterium slows channel opening.
The time-course of H+ or D+ current activation during depolarizing pulses was fitted by a single exponential after a delay to obtain τact, as was shown in the inset in Fig. 5. Mean values for τact at various pD (solid symbols) and pH (open symbols) are plotted in Fig. 12, all for ΔpL = 0. It is unclear from these data whether there might be some effect of the absolute value of pL on τact. However, all the mean τact values in D2O are slower at each potential than any of the values in H2O. The average of the ratios at all potentials ≥60 mV of the mean τact values in D2O to H2O at 0.5 U lower pLi was 3.21 at pD 8, 3.19 at pD 7, and 2.96 at pD 6. In summary, D2O slows τact by about threefold.
Because there was substantial variability of τact from one cell to another, comparisons were also made in individual cells at effectively symmetrical pH or pD. The average ratio of τact in D2O to that in H2O plotted in Fig. 13 reveals that τact was 2.0–3.6 times slower in D2O. The slowing was not noticeably voltage dependent. There is a suggestion that the slowing effect was greater at higher pD (or pH). If the ratios at all voltages in each solution are averaged, the slowing effect was 2.17 at pD 6 compared with pH 5.5, 3.06 at pD 7 compared with pH 6.5, and 3.21 at pD 8 compared with pH 7.5. The solid symbols include only cells studied with D2O pipette solutions, the open squares show data from cells with H2O in the pipette. The slowing of τact by D2O appears to be attenuated in these cells, possibly reflecting the small amount of H2O inside the cell, although the difference is not significant. In summary, D2O slows τact about threefold, and this effect appears to be voltage independent.
Deuterium does not strongly affect deactivation kinetics.
The channel closing rate was examined by fitting the time course of the decay of tail currents (materials and methods), such as those illustrated in Fig. 1, A–C. The average values of τtail obtained in effectively symmetrical solutions are plotted in Fig. 14. There is a suggestion in the data that τtail was slightly slower at higher pL, and in D2O compared with H2O. The average ratios at all potentials of the mean τtail data for essentially symmetrical pL are 1.31 (pD 8/pH 7.5), 1.04 (pH 7.5/pD 7), 1.23 (pD 7/pH 6.5), 1.05 (pH 6.5/pD 6), and 1.51 (pD 6/ pH 5.5). The apparent slowing by D2O was thus 23– 51%, and some part of this effect may be ascribable to increasing pLi.
In some cells τtail is independent of pHo (DeCoursey and Cherny, 1996a; Cherny et al., 1997), but the effects of pHi have not been clearly determined. Therefore, we attempted to compare τtail in H2O and D2O at similar pLi in the same cell by varying the NH4+ gradient. Increasing pHi in individual cells at constant pHo consistently slowed τtail by a small amount (not shown). When D2O was compared with H2O at a constant NH4+ gradient, i.e., at nearly constant pLi (see above), there was also a consistent slowing of τtail in nearly every cell, by roughly 50%, consistent with the average values given above.
Deuterium effects in cell-attached patches.
Fig. 15 illustrates putative H+ currents in a cell-attached patch. The cell was bathed with isotonic KMeSO3 solution to depolarize the membrane to near 0 mV. During depolarizations positive to 0 mV, there are slowly activating outward currents that resemble H+ currents (cf. DeCoursey and Cherny, 1995), as well as brief discrete openings of some other channel(s). When H2O in the bath was replaced with D2O, the outward currents became much smaller and appeared to activate even more slowly. This isotope effect is comparable to the effects seen in whole-cell configuration, but larger than reported for other ion channels (Table III). Therefore, we conclude that the slowly activating outward currents were in fact H+ currents.
Absolute H+ or D+ permeability of the cell membrane (not through proton channels).
The “leak” current at subthreshold voltages usually decreased when D2O replaced H2O. However, it appears extremely unlikely that the leak is carried primarily by H+ or D+. Attempts to calculate the H+ permeability, PH,, of the leak current using the Goldman-Hodgkin-Katz (GHK) current equation (Goldman, 1943; Hodgkin and Katz, 1949):
where IH and PH are expressed normalized to membrane area estimated assuming that the specific capacitance is 1 μF/cm2, revealed numerous inconsistencies with this idea. The slope conductance of leak currents (defined as time-independent currents at subthreshold potentials) rarely changed by more than twofold/U change in pH or pD, and not always in the same direction. For a large pL gradient (e.g., pD 8//6), leak currents at negative potentials but positive to EL were inward, giving a negative calculated PL. Calculated values for PL decreased substantially at low pLo, even when the observed leak slope conductance was increased. Finally, the apparent reversal potential of the leak current, which was not well defined because the leak currents were often small, was usually closer to 0 mV than to E L, and did not always change in the “right” direction when pLo was varied. In summary, there is no evidence that H+ carries a significant fraction of the leak current. An upper limit on the passive membrane permeability to H+ or D+ can be given as <<10−4 cm/s at pHi 5.5 or pDi 6. By comparison, when the gH is fully activated, PH exceeds 1 cm/s at pH 8.0//7.5 (calculated from data in Cherny et al., 1995).
The deuterium isotope effects observed provide information about H+ permeation as well as the regulation of gating by protons (or deuterons). The main results are: (a) D+ permeates proton channels. (b) The relative permeability of proton channels is >108 greater for D+ than for TMA+. (c) The H+ conductance through proton channels is ∼1.9 times that of D+. (d) D+ regulates the voltage dependence of H+ channel gating much like H+. (e) The threshold for activating the proton conductance is a linear function of Vrev and changes 40 mV/U change in ΔpH or ΔpD. (f) D+ currents activate with depolarization ∼3 times slower than H+ currents, but deactivation is at most 1.5-fold slower in D2O. (g) At least 5.4 equivalent gating charges move across the membrane field during proton channel opening in D2O and in H2O. (h) The upper limit of any proton leak conductance of the membrane of rat alveolar epithelial cells must be <<10−4 cm/s. When the gH is fully activated, PH exceeds 1 cm/s.
Properties of Proton Channels
Proton channels are extremely selective.
At high pD, the D+ permeability was >108 greater than the TMA+ permeability. The calculated permeability ratio PTMA/PD decreased as pD increased, by about 10-fold/U change in pDi. Although a concentration dependent permeability ratio cannot be ruled out, it seems more reasonable to suppose that deviations of Vrev from ED are due to imperfect control of pD, rather than to finite permeability of the channel to other ions. Several other H+ channels have been reported to have comparably high selectivity for H+, including the F0 component of H+-ATPase (Althoff et al., 1989; Junge, 1989) and the M2 viral envelope protein (Chizhmakov et al., 1996).
Protons rather than hydroxide ions carry the current.
The substantially lower conductance of proton channels in D2O than in H2O suggests that the charge-carrying species is H+ (or D+) rather than OH− (or OD−). The isotope effect for D+ is large because its mass is twice that of H+, but OD− is only 6% heavier than OH−, and thus a much smaller isotope effect is to be expected: 41% for D+ vs. 3% for OD− for a classical square-root dependence on the mass of reactants (Glasstone et al., 1941). A similar argument can be made against H3O+ which would have a predicted isotope effect of just 8% over D3O+. However, the extremely high selectivity of the gH has been ascribed to a Grotthuss-type or proton-wire permeation mechanism, which could exist for L+ or OL−, but not L3O+ (Nagle and Morowitz, 1978; DeCoursey and Cherny, 1994). Additional evidence supporting H+ rather than OH− as the charge carrying species is that the gH increases ∼1.7-fold/U decrease in pHi over the range pHi 7.5–4.0 (DeCoursey and Cherny, 1995, 1996a), i.e., as [H+]i increases and [OH−]i decreases and [OH−]o remains constant. Finally, the reduction of outward current in cell-attached patches when the bath solvent is changed from H2O to D2O (Fig. 15), is consistent with L+ efflux across the membrane from the cell to the pipette, but not OL− influx from the pipette into the cell.
Voltage-gated H+ and D+ currents pass through channels, not the phospholipid bilayer membrane: the gH is not a membrane leak.
The finding that the voltage-activated and time-dependent H+ conductance is clearly larger than the D+ conductance provides further support for the idea that this conductance occurs through specialized membrane transporters, presumably proteins, and not simply through leaks in the bilayer. The conductance of phospholipid bilayers to D+ is similar to that of H+ (Perkins and Cafiso, 1986; Deamer, 1987; Gutknecht, 1987). The proton (or OH−) permeability, PH, of lipid bilayer membranes is several orders of magnitude higher than its permeability to other cations. Reported values for PH vary widely, from 10−9 to <10−3 cm/s in lipid bilayers and from 10−5 to 10−3 cm/s in biological membranes (reviewed by Deamer and Nichols, 1985). At least part of this variability is due to a dependence on the nature of the membrane and the pH gradient, ΔpH (Perkins and Cafiso, 1986)—at ΔpH = 1.0 in membranes of varying lipid composition, PH ranged from 2.0 × 10−7 to 1.8 × 10−5 cm/s (Perkins and Cafiso, 1986). We suspect that no more than a very small fraction of our leak current at subthreshold potentials is carried by H+ or D+. This leak current provides an upper limit of PH <<10−4 cm/s in rat alveolar epithelial cells, providing no indication of any unusual H+ permeability of these particular biological membranes. Even if the leak were carried entirely by H+ or D+, PH increases by 3–4 orders of magnitude during depolarization from subthreshold to large positive potentials. It is difficult to imagine that a transient water-wire spanning the membrane would exhibit consistent, well- defined voltage- and time-dependent gating.
If we convert the observed voltage-gated H+ current to permeability, PH, using the GHK current equation (Goldman, 1943; Hodgkin and Katz, 1949), PH increases with depolarization approaching a limiting value at any given ΔpH. However, the value calculated for PH is much larger at high pHi, because of the relative insensitivity of the H+ conductance, gH, to absolute pH (Cherny et al., 1995; DeCoursey and Cherny, 1995). The limiting value for PH is about 1.1 × 100 cm/s at pH 8.0//7.5, 1.7 × 10−1 cm/s at pH 7.0//6.5, and 1.4 × 10−2 cm/s at pH 6.0//5.5 (recalculated from data in Cherny et al., 1995). Clearly, the GHK formalism is not a useful means of expressing PH through the voltage- activated gH, because its value is nowhere near being concentration-independent. That the PH values obtained for the voltage-gated gH are 3–9 orders of magnitude greater than those for H+/OH− conductivity through lipid bilayers makes it clear that the voltage- activated gH requires a special transport molecule and cannot reasonably be ascribed to H+ permeation through the phospholipid component of the cell membrane.
What is the rate-limiting step in H+ permeation?
The ratio of H+ current to D+ current was 1.65, 1.91, and 1.92 at pD 6, 7, and 8, respectively. Nearly all the H+ that carry current during a depolarizing pulse are derived from buffer molecules that were protonated before the pulse (DeCoursey, 1991). If diffusion of protonated buffer to the channel were rate limiting, one would predict a smaller isotope effect on the conductance. Protonated or deuterated buffer should have almost identical diffusion coefficients. However, the 25% greater viscosity of D2O than H2O (Table I) would impede the diffusion of buffer molecules. That the gH is reduced by almost 50% in D2O is inconsistent with buffer diffusion being rate determining. We have shown recently that above 10 mM buffer there is negligible limitation of H+ current by the diffusion of buffer at either side of the membrane (DeCoursey and Cherny, 1996b). In contrast, the smaller deuterium isotope effect on the conductance of most ion channels is consistent with diffusion of permeant ions being the rate-determining factor (Table III).
If H+ permeation were set by the hydrodynamic mobility of H3O+, then the H+/D+ conductance ratio should similarly correspond with the relative viscosities and dielectric constants of H2O and D2O (Lengyel and Conway, 1983). In fact, the relative mobility of H+ in H2O to D+ in D2O is significantly larger, namely 1.41 compared with 1.17 for KCl in H2O vs. D2O at 20°C (interpolated from the data of Lewis and Doody, 1933), indicating that a more rapid transfer mechanism for H+ exists, namely the “Grotthuss” mechanism in which protons hop from one water molecule to another. An isotope effect of 1.4–1.5 might therefore be expected if H+ or D+ conduction to the mouth of the pore were rate determining, or if permeation through the channel involved a mechanism like H+ or D+ diffusion in bulk water. Indeed, the relative conductance of H+ to D+ through gramicidin is of this magnitude, 1.34 at 10 mM L3O+, consistent with the approach of L3O+ to the channel being rate limiting, and 1.35 at 5 M L3O+ where the gramicidin channel current is saturated and the ratio presumably reflects that of permeation mechanism (Akeson and Deamer, 1991). The gH/gD ratio in voltage-gated H+ channels was larger than can be accounted for by diffusion of either buffer or L3O+ molecules, strongly suggesting that the rate-determining step in permeation occurs in the channel itself. Furthermore, the larger isotope effect in voltage-gated channels than in gramicidin suggests that H+ permeates by a different mechanism than gramicidin, in which H+ is believed to hop across a continuous hydrogen-bonded chain of water molecules filling the pore (Myers and Haydon, 1972; Levitt et al., 1978; Finkelstein and Andersen, 1981; Akeson and Deamer, 1991). Perhaps voltage-gated H+ channels are not simple water-filled pores, but include amino acid side groups in the hydrogen-bonded chain, as proposed previously to account for their high selectivity and nearly pH-independent conductance (DeCoursey and Cherny, 1994, 1995; Cherny et al., 1995), by analogy with the proton wire mechanism proposed by Nagle and Morowitz (1978) to explain H+ transport through the “proton channel” component of mitochondrial and chloroplast H+-ATPases and bacteriorhodopsin. In summary, although the permeation of H+ through gramicidin behaves in a manner consistent with the behavior of H+ in bulk water solution, the permeation of H+ through voltage-gated channels appear to behave differently.
To explain the apparent pH independence of the H+ conductance of bilayer membranes, Nagle (1987) suggested that the rate-determining step might be the breaking of hydrogen bonds between water molecules. Applied to H+ channel currents, the H+ conductance might have an activation energy like that of hydrogen bond cleavage. The isotope effect for cleavage of an ordinary hydrogen bond in liquid water is ∼1.4 (Walrafen et al., 1996). The observed ratio of H+ to D+ current, ∼1.65–1.92, is significantly larger, suggesting that the rate determining step resides elsewhere. If a quantum-mechanical tunnel transfer within the pore were rate determining, then a much larger isotope effect would be expected, for example, 6.1 calculated for the relative mobilities calculated for tunnel transfers in water (Conway et al., 1956). Although H+ tunneling may occur in the channel, it evidently is not rate limiting.
As discussed above, we imagine that the H+ channel is not a water-filled pore but is most likely composed of some combination of amino acid side groups and water molecules linked together in a membrane-spanning hydrogen-bonded chain. Proton conduction is believed to occur by a Grotthuss or proton wire mechanism, which requires both hopping and reorientation steps (see introduction; Nagle and Morowitz, 1978; Nagle and Tristram-Nagle, 1983). By analogy with ice, the mobility of the H+ “ionic defect” is 6.4 × 10−3 cm2 V−1 s−1 (at −5°C, Kunst and Warman, 1980), about an order of magnitude greater than the Bjerrum L defect mobility, 5 × 10−4 cm2 V−1 s−1 (at 0°C, Camplin et al., 1978), suggesting that the turning step may be rate determining. However, proton transfer may be slower when it occurs between two dissimilar elements of the hydrogen-bonded chain. For example, proton transfer is slowed in mixed solvents because protons become effectively trapped by the solvent molecule with higher H+ affinity (Lengyel and Conway, 1983). It is intriguing that the mobility of H+ in ice exhibits a large isotope effect, 2.7 for H+/D+ at −5°C (Kunst and Warman, 1980). Furthermore, the reorientation of hydrogen bonds during proton transport in ice exhibits a H2O/D2O ratio of ∼1.6 (at −10°C, Eigen et al., 1964), suggesting by analogy that the turning step for water which is constrained in a channel pore may exhibit a larger isotope effect than water in free solution. Although the rate-limiting step in H+ permeation appears to occur within the conduction pathway, we cannot resolve whether the hopping or turning step is rate determining.
Are H+ channels really ion channels?
In Table IV deuterium isotope effects on various membrane transporters other than channels are listed. The precise values depend strongly on the conditions of the measurement, but in general it appears that more complex transport mechanisms exhibit stronger isotope effects on transport rates, >1.7, compared with <1.5 for ion channel permeation (Table III). This result strengthens the conclusion that the H+ channel is not a simple water-filled pore, which was based on its high H+ selectivity and nearly pH independent conductance. If voltage-gated H+ channels are not water-filled pores, should they be considered ion channels at all? H+ current does not require ATP or any counter-ion, so the only possibly more accurate term would be a carrier. The essential difference between a carrier and a channel is that each ion transported through a carrier requires a conformational change in the molecule which changes the accessibility of the ion from one side of the membrane to the other, whereas an open channel conducts ions without obligatory conformational changes. (Of course, there are significant interactions between conducted ions and the channel pore.) Biological channels also exhibit gating, without which they would simply be holes in the membrane. The voltage-gated H+ channel exhibits well-defined time-, voltage-, and pH-dependent gating. That the conduction process involves protons hopping across a hydrogen-bonded chain seems a minor distinction. The two-stage hop-turn mechanism of the proton-wire (Nagle and Morowitz, 1978) could perhaps be described technically as alternating-access, in that the hydrogen-bonded chain must re-load after each H+ conduction event. However, a hop-turn mechanism is also believed to occur when H+ are conducted through gramicidin, in which the proton wire is composed entirely of water molecules, and there seems to be consensus that gramicidin is an ion channel, not a carrier. On balance, we prefer the term channel, but recognize that H+ conduction by a proton wire (hydrogen-bonded chain) mechanism may bear some similarities to the alternating access mechanism which defines carriers and that H+ channels may be unique among ion channels in not having a water-filled pore.
Deuterium Isotope Effects on Gating
Regulation of H+ channel gating by pH.
The rates of H+ channel opening (activation) and closing (deactivation) are voltage dependent, both processes becoming faster at large voltages. Byerly et al. (1984) found that increasing pHi or lowering pHo shifted the voltage dependence of activation kinetics of H+ currents in snail neurons to more positive potentials but that lowering pHo slowed activation more than could be explained by a simple voltage shift. Subsequent studies in a variety of cells leave the impression that both low pHo and high pHi slow activation somewhat more than expected for a simple voltage shift (Kapus et al., 1993; Cherny et al., 1995; DeCoursey and Cherny, 1996a), although in some cases a simple shift by pHo was observed (Barish and Baud, 1984; DeCoursey and Cherny, 1995). Studied in inside-out membrane patches, increasing pHi slowed activation by approximately fivefold/U in addition to shifting the voltage dependence of channel opening (DeCoursey and Cherny, 1995). The effects of pH on deactivation are substantially weaker than on activation. The voltage dependence of τtail was shifted at most 20 mV/U change in ΔpH in alveolar epithelial cells (Cherny et al., 1995). In THP-1 monocytes changing pHo by 2 U had no detectable effect on τtail (DeCoursey and Cherny, 1996a). Here we report that H+ current activation is slowed dramatically in D2O whereas deactivation was barely affected.
Deuterium isotope effects on other channels.
Deuterium isotope effects on several voltage-gated ion channels are summarized in Table III. Two features are noteworthy. Deuterium slows the opening rate of all channels studied, but the slowing is much greater for H+ channels. For Na+ or K+ channels, τact is slowed only ∼1.4-fold near 0°C, and this effect is halved at 10–14°C (∼1.2-fold slowing) and undetectable 15–20°C (Schauf and Bullock, 1982; Alicata et al., 1990). The relatively subtle effects on τact of other channels have been ascribed to changing solvent structure (e.g., Schauf and Bullock, 1980, 1982). The effect on Na+ channel inactivation is significantly larger, decreases at higher temperatures, and may reflect a different mechanism. Also remarkable is the solvent-insensitivity of deactivation, a result that appears to hold also for voltage-gated H+ channels. It is conceivable that the greater deuterium sensitivity of activation than deactivation reflects some common principle of the mechanism of ion channel gating. However, the large isotope effect on H+ channel activation seems to implicate a protonation/deprotonation reaction in gating, rather than a mechanism involving changes in solvent structure.
Deuterium isotope effects on H+ channels.
The opening rate of H+ channels was 3.2, 3.1, and 2.2 times slower in D2O at pD 8, pD 7, and pD 6, respectively. In contrast, the closing rate was slowed only 1.5-fold or less. In the model proposed to account for the regulation of the voltage dependence of gating by pH, the first step in channel opening is deprotonation at an externally accessible site on the channel, and the first step in channel closing is deprotonation at an internally accessible site (Cherny et al., 1995). If deprotonation at the external site were the rate-determining step in channel opening, then the observed slowing of τact could reflect an increase in the pKa of this site in D2O by 0.34–0.51 U. We give more weight to the larger D2O effects, because factors such as H2O contamination and the possibility that other deuterium-insensitive steps in gating may contribute to the observed kinetics would tend to diminish the size of the observed effect. We conclude that the pKa of the external site most likely increases by ∼0.5 U in D2O. The pKa of simple carboxylic and ammonium acids increases in D2O by ∼0.5–0.6 U, whereas the pKa of sulfhydryl acids increases only 0.1–0.3 U (Schowen, 1977). The observed slowing of τact thus speaks against cysteine as the amino acid comprising the hypothetical site. We conclude that the modulatory site that governs the opening of H+ channels is most likely a histidine, lysine, or tyrosine residue. The stronger D2O isotope effect on activation than deactivation suggests that either the external and internal regulatory sites are chemically different, or the first step in channel closing occurs before deprotonation at the internal site.
One remarkable aspect of the data in Fig. 11 is that there is no suggestion of saturation of the relationship between Vrev and Vthreshold. We previously reported saturation of the shift in the position of the gH-V relationship above pHo 8, with only a 10–20-mV shift between pHo 8 and pHo 9 (Cherny et al., 1995). In the present study, similar apparent saturation was observed, and extending the measurement to pHo 10 resulted in no further shift relative to pHo 9. However, we found that at high pHo, Vrev deviated substantially from EH. In the previous study we felt that we could not resolve Vrev at pHo 9 due to the rapid kinetics. Although tail currents at pHo 9 or pHo 10 were resolved less well than at lower pHo, when we plot Vthreshold against the best estimate of Vrev (Fig. 11), the data fall on the linear relationship consistent with the other, better determined data points. It appears that there is an anomalous loss of control over pHi at very high pHo. It is difficult to imagine that pHo is not well established by 100 mM buffer in the bath, and, assuming that Vrev reflects the true ΔpH, pHi must increase a full unit when pHo is changed from 9 to 10. One possibility is that some additional pH-regulating membrane transport process is working under these conditions. For example, a recently described Cl−/OH− exchanger (Sun et al., 1996) working “backwards” might exchange external OH− for internal Cl−, in spite of the rather low (4 mM) Cl− concentration in the pipette solutions. Although we cannot explain the mechanism, the phenomenon merits further study. The lack of saturation complicates estimation of the pKa of the putative regulatory protonation sites on H+ channels.
Predicting the voltage dependence of the gH in intact cells.
The definition of Vthreshold is certainly arbitrary, because by using longer pulses, heavier filtering, and higher gain, it is possible to detect smaller and smaller currents, and ultimately Vthreshold has no precise theoretical meaning. Nevertheless, predicting the circumstances under which the gH might be activated in vivo is facilitated by some estimate of Vthreshold. The slope of the line in Fig. 11 for the H2O data corresponds with a 40.0-mV shift/U change in ΔpH, if Vrev changes by 52.4 mV/U ΔpH, as reported previously (Cherny et al., 1995), or a 44.4 mV/U shift if Vrev changed according to EH. The slope in D2O was virtually identical. Thus the previous conclusion that the voltage-activation curve is shifted by ∼40 mV/U change in ΔpH is in excellent agreement with the present data both in H2O and in D2O. We previously proposed that Vthreshold in intact cells could be predicted from the empirical relationship:
where V0 was typically 20 mV, but varied substantially from cell to cell (Cherny et al., 1995). This relationship is based on the nominal ΔpH. Considering the remarkably linear relationship in Fig. 11 between Vthreshold and Vrev, we suggest that a more accurate prediction can be based of the true ΔpH, which we feel is reflected more closely by the observed Vrev than by the applied ΔpH. The new, improved relationship (in H2O) is:
This relationship is very similar to that described by Eq. 6, in predicting a ∼40-mV shift in Vthreshold/U change in ΔpH, and Vthreshold near +20 mV at symmetrical pH (ΔpH = 0), but emphasizes the use of Vrev as the ultimate indication of the true ΔpH. The dotted reference line in Fig. 11 illustrates that Vthreshold is positive to Vrev over the entire voltage range studied. The regulation of the voltage-activation curve by ΔpH thus results in only steady-state outward currents throughout the physiological range.
We are grateful for constructive comments on the manuscript by Peter S. Pennefather, Duan Pin Chen, the reviewers, and Noam Agmon, who also generously provided preprints. The authors appreciate the excellent technical assistance of Donald R. Anderson, and thank Charles Butler for some determinations of the pKa of buffers in normal and heavy water.
This study was supported by a Grant-in-Aid from the American Heart Association and by National Institutes of Health Research Grant HL-52671 to T. DeCoursey.
Preliminary accounts of this work have been previously reported in abstract form (Cherny, V.V., and T.E. DeCoursey. 1997. Biophys. J. 72: A266; DeCoursey, T.E., and V.V. Cherny. 1997. Biophys. J. 72:A108).
Abbreviations used in this paper: ΔpD, pD gradient (pDo − pDi); ΔpH, pH gradient (pHo − pHi); ED, Nernst potential for D+; eff, effective composition of the intracellular solution given the assumptions discussed in Strategic Considerations; EH, Nernst potential for H+; EL, either EH or ED; gD, D+ conductance; gH, H+ conductance; GHK, Goldman-Hodgkin-Katz; L+, either H+ or D+; L3O+, either H3O+ or D3O+; pD, the equivalent in D2O of pH in water; Pd, diffusional water permeability; PD, permeability to D+; Pf, osmotic water permeability; PH, permeability to H+ calculated with the GHK voltage equation; pHi, intracellular pH; pHo, extracellular pH; pL, either pH or pD; Posm, water permeability; PTMA, permeability to TMA+; OL−, either OH− or OD−; τact, time constant of activation; τtail, tail current or deactivation time constant; TMA+, tetramethylammonium; TMAMeSO3, tetramethylammonium methanesulfonate; Vhold, holding potential; Vrev, reversal potential; Vthreshold, threshold potential for activating proton currents.
Address correspondence to Dr. Thomas E. DeCoursey, Department of Molecular Biophysics and Physiology, Rush Presbyterian St. Luke's Medical Center, 1653 West Congress Parkway, Chicago, IL 60612. Fax: 312-942-8711; E-mail: email@example.com