From the Department of Molecular Biophysics and Physiology, Rush Presbyterian St. Luke's Medical Center, Chicago, Illinois 60612
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 (
act 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 (
tail) 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
).
Table I. Properties of H2O and D2O at 20°C |
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 (0) 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
pH, within a large pH range encompassing
physiological values. Thus the position of the voltage
activation curve can be predicted from the pH gradient,
pH, 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.
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.
Solutions
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:
![]() |
(1) |
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.
pD Measurement
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.
Table II. pKa of Buffers in H2O and D2O |
Electrophysiology
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 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 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.
Data Analysis
The time constant of H+ current activation, act, was obtained by
fitting the current record by eye with a single exponential after a
brief delay (as described in DeCoursey and Cherny, 1995
):
![]() |
(2) |
where I0 is the initial amplitude of the current after the voltage
step, I is the steady-state current amplitude, t is the time after the
voltage step, and tdelay is the delay. The H+ current amplitude is
(I0 I
). No other time-dependent conductances were observed
consistently under the ionic conditions employed. Tail current
time constants,
tail, were fitted either to a single decaying exponential:
![]() |
(3) |
where I0 is the amplitude of the decaying part of the tail current, or to the sum of two exponentials:
![]() |
(4) |
where An are amplitudes and n are time constants.
Conventions
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.
Strategic Considerations
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, pH, 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
pH 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.
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.
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 104 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.
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.
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. pH 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
pH 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 resultVrev 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, pD, 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
pD = 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, act, in the same cell at pDo 8, 7, and 6 are
plotted in Fig. 5. At each pDo
act is clearly voltage dependent, decreasing with depolarization. Lowering pDo
appears to shift the
act-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
act
varied from cell to cell, the effects of changes in pDo in
each cell were quite similar to those illustrated.
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
Families of currents in the same cell in H2O and D2O
are illustrated in Fig. 8. To keep pL approximately
constant, we compared pHo 6.5//pHi,eff 6.5 and pDo 7//
pDi 7 (Fig. 8, A and B, respectively). In D2O the currents are smaller and activate more slowly.
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 pD just as in
H2O. However, the absolute voltage dependence might
be different in the two solvents. To address this possibility we compared similar
pH and
pD 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 102 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 pD as by
pH.
The time-course of H+
or D+ current activation during depolarizing pulses was
fitted by a single exponential after a delay to obtain act,
as was shown in the inset in Fig. 5. Mean values for
act
at various pD (solid symbols) and pH (open symbols) are plotted in Fig. 12, all for
pL = 0. It is unclear from
these data whether there might be some effect of the
absolute value of pL on
act. However, all the mean
act
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
act 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
act by about threefold.
Because there was substantial variability of act from
one cell to another, comparisons were also made in individual cells at effectively symmetrical pH or pD. The
average ratio of
act in D2O to that in H2O plotted in
Fig. 13 reveals that
act 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
act 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
act about threefold, and this effect appears to be voltage independent.
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 tail obtained in effectively symmetrical
solutions are plotted in Fig. 14. There is a suggestion in
the data that
tail was slightly slower at higher pL, and in
D2O compared with H2O. The average ratios at all potentials of the mean
tail 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 tail 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
tail 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
tail 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
tail in nearly every cell,
by roughly 50%, consistent with the average values
given above.
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.
Table III. Deuterium Isotope Effects on Other Channels (temperature, °C) |
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
):
![]() |
(5) |
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 <<104 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 pH or
pD. (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
).
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.
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,
pH
(Perkins and Cafiso, 1986
)
at
pH = 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
pH. 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.
Deuterium Permeation
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.
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.
Table IV. Comparison of D+ and H+ Flux through Other Membrane H+ Transporters |
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
tail was shifted at
most 20 mV/U change in
pH in alveolar epithelial
cells (Cherny et al., 1995
). In THP-1 monocytes changing pHo by 2 U had no detectable effect on
tail (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 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, act 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
act 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.
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
act 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
act 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
pH, 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.
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 pH, if Vrev changes by 52.4 mV/U
pH, 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
pH 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:
![]() |
(6) |
where V0 was typically 20 mV, but varied substantially
from cell to cell (Cherny et al., 1995). This relationship
is based on the nominal
pH. 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
pH, which we feel is reflected more
closely by the observed Vrev than by the applied
pH.
The new, improved relationship (in H2O) is:
![]() |
(7) |
This relationship is very similar to that described by Eq. 6, in predicting a ~40-mV shift in Vthreshold/U change in
pH, and Vthreshold near +20 mV at symmetrical pH
(
pH = 0), but emphasizes the use of Vrev as the ultimate indication of the true
pH. 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
pH thus results in only
steady-state outward currents throughout the physiological range.
Original version received 27 November 1996 and accepted version received 27 January 1997.
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: tdecours{at}rpslmc.edu
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).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.