Ethics statement

Surgical removal of ovary tissue from adult Xenopus laevis females followed registered protocols approved by the relevant state authority (Landesamt für Gesundheit und Soziales Berlin, Reg. No. O 0308/06) and the local ethics committee (Tierschutzbeirat), in strict accordance with the German Animal Protection Act (Tierschutzgesetz). Animals were anesthetized by immersion in water containing 0.2% w/v tricaine (MS-222, Sigma, Deisenhofen, Germany) for 5 min, and subsequently placed on ice during surgical treatment. All efforts were made to minimize animal suffering.

Protein expression in Xenopus oocytes

Xenopus oocytes were obtained by collagenase treatment after partial ovariectomy from Xenopus laevis females. cRNAs were prepared using the SP6 mMessage mMachine Kit (Agilent Technologies, Santa Clara, CA). A 50 nl aliquot containing 20–25 ng rat gastric H,K-ATPase α-subunit cRNA and 5 ng wild-type H,K-ATPase β-subunit cRNA was injected into each cell. The variant HKαS806C, which carries the mutation S806C within the M5/M6 loop to enable site-specific labeling with TMRM (see Figure S2), was used as parent construct for mutagenesis and is termed “wild-type” herein. The S806C mutation does not affect ion transport activity [32], [33]. Mutagenesis was performed by recombinant PCR and verified by DNA sequencing (Eurofins MWG Operon, Ebersberg, Germany). After cRNA injection, oocytes were kept in ORI buffer (110 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 5 mM HEPES, pH 7.4, plus 50 mg/l gentamycin) at 18°C for two days.

Experimental solutions

Experimental solutions were Na buffer, TMA buffer, NMDG buffer, (20 mM TEACl, 5 mM BaCl₂, 5 mM NiCl₂ and 90 mM NaCl, TMA-Cl or NMDG-Cl, respectively). For measurements at extracellular pH (pH_ex) of 7.4, the solution was buffered with 10 mM HEPES, whereas 10 mM MES was used for experiments at pH_ex 5.5. The pH of the buffers used is indicated by a lower index. For measurements in presence of extracellular K⁺, 5 mM NaCl of the Na_7.4 buffer (or Na_5.5 buffer) were replaced by 5 mM K⁺. For intracellular acidification measurements in Na_7.4 buffer, 40 mM NaCl were replaced by 40 mM Na-butyrate. At pH_ex 7.4, a concentration of 40 mM butyrate corresponds to 100 µM undissociated (thus membrane-permeable) butyric acid, which was reported to increase cytosolic [H⁺] of oocytes from 50 nM to 160 nM (corresponding to an intracellular pH change from ∼7.3 to ∼6.8 [35]). All solutions contained 100 µM ouabain to inhibit the endogenous Xenopus Na,K-ATPase.

Voltage clamp fluorometry

Site-specific labeling of H,K-ATPase α-subunit mutant S806C (HKαS806C) upon expression oocytes was achieved by incubating oocytes in Na_7.4 buffer with 5 µM TMRM (tetramethylrhodamine-6-maleimide, Molecular Probes) for 5 min at room temperature in the dark, followed by extensive washes in dye-free Na_7.4 buffer. Labeled oocytes were transferred into an oocyte perfusion chamber (model RC-10, Warner Instr., Hamden, CT), which was mounted on the stage of an epifluorescence microscope (Axioskop 2FS; Carl Zeiss, Göttingen, Germany) equipped with a 40× water immersion objective (numerical aperture = 0.8). Fluorescence was excited with a 100 W tungsten lamp using a 535DF50 excitation filter, a 565 EFLP emission filter and a 570DRLP dichroic mirror (Omega Optical, Battleborough, USA). Fluorescence monitoring used a PIN-022A photodiode (United Detector Technologies, Torrence, CA) mounted to the microscope camera port, whose photocurrents were amplified by a DLPCA-200 low-noise current amplifier (FEMTO Messtechnik GmbH, Berlin, Germany). Control of transmembrane voltage was achieved by means of a Turbotec 05 two-electrode voltage clamp amplifier (npi, Tamm, Germany). Fluorescence and current signals were recorded simultaneously using a Digidata 1322A interface and pClamp 9.2 software (Molecular Devices, Sunnyvale, CA).

Rb⁺ uptake assay

Two days after injection, non-injected control oocytes and H,K-ATPase-expressing oocytes were preincubated for 15 min in TMA_7.4 buffer, containing 100 µM ouabain for complete inhibition of the endogenous Na,K-ATPase. Oocytes were then incubated for 15 min in Rb⁺-flux-buffer (5 mM RbCl, 85 mM TMACl (NMGCl or NaCl), 20 mM TEACl, 5 mM BaCl₂, 5 mM NiCl₂, 10 mM MES, pH 5.5 or pH 7.4, 100 µM ouabain). For intracellular acidification measurements in Na_7.4 buffer, 40 mM NaCl in the Rb⁺-flux-buffer were replaced by 40 mM Na-butyrate. In NMDG- or TEA-based solutions, 40 mM NMDG (or TEA) were substituted by 40 mM butyric acid prior to pH adjustment using HCl. Temperature-dependent Rb⁺ uptake measurements (between 18°C and 34°C) were performed by incubation in an HLC thermomixer (Ditabis, Pforzheim, Germany). Rb⁺ uptake under voltage control was measured using the aforementioned two-electrode voltage clamp setup to apply −100 mV membrane potential during incubation in Rb⁺-flux-buffer.

After three washing steps in Rb⁺-free TMA_7.4 buffer and one wash in Millipore water, each individual oocyte was homogenized in 1 ml of Millipore water. 20 µl samples of the oocyte homogenates were automatically transferred into the transversely heated graphite furnace of an AAnalyst800™ atomic absorption spectrometer (Perkin Elmer, Waltham, MA). After two drying steps at 110°C and 130°C and a pyrolysis step at 500–600°C, atomization was carried out at 1700–1800°C. After each measurement, the graphite furnace was heated to 2400°C for cleaning. Rubidium absorption was measured at 780 nm using a Rubidium hollow cathode lamp (Photron, Melbourne, Australia). After Zeeman-background correction, Rb⁺ contents were calculated from the integrated peak area of the signal according to a standard calibration curve. Between 0 and 70 µg/L RbCl an excellent linearity (r²≥0.99) was observed. The detection limit of Rb⁺ was in the upper picomolar range (characteristic mass: 10 pg).

Effect of extra- and intracellular pH on presteady-state fluorescence changes

The aim of this study was to investigate electrogenic partial reactions within the H⁺ translocating branch of gastric H,K-ATPase and to scrutinize, whether and how the concept of high-field access channels established for Na,K-ATPase [12] can be transferred to the H⁺ pump. For that purpose, we performed voltage clamp fluorometry on gastric H,K-ATPase mutant S806C under various ionic conditions. Figure 2A shows typical fluorescence signals at pH_ex 7.4 resulting from voltage pulses to a series of test potentials between −180 and +60 mV, which were recorded in the absence of K⁺ and with 90 mM Na⁺ in the extracellular solution. The fluorescence of the dye attached to the extracellular end of helix M5 (see structural model in Figure S2), increases upon jumps to negative potentials and decreases at depolarizing membrane voltage. In analogy to Na,K-ATPase, positive voltages should favor the transition to E₂P, whereas negative voltage steps drive the enzyme into E₁P. According to the crystal structures of several reaction intermediates of the related SERCA Ca²⁺-ATPase [36], [37], [38], [39], the central helix M5 moves in relation to the surrounding helices during the cycle. Since TMRM is sensitive to hydrophobicity and collisional quenching, the observed fluorescence signals presumably result from a motion of the extracellular end of M5 from a buried, sheltered environment in E₁P into a more aqueous, quenching environment in E₂P.

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Figure 2. Effects of extracellular pH on the E₁P/E₂P distribution of gastric H,K-ATPase.

(A,B) Fluorescence responses of site-specifically labeled gastric H,K-ATPase under K⁺-free conditions (90 mM Na⁺ in the extracellular solution) upon voltage jumps from −40 mV to voltages between −180 mV and +60 mV in (−20 mV steps, see inset in A) at an extracellular pH of 7.4 (A) or 5.5 (B). (C) Voltage dependence of normalized fluorescence amplitudes (1-ΔF/F) from experiments as in (A,B) for pH_ex 5.5 (○), and for pH_ex 7.4 (•). Data are means±S.E. of 11–14 oocytes. Superimposed as dashed lines are curves resulting from fits of a Boltzmann-type function to the data sets (pH_ex 7.4: V_0.5 = −19.7±5.4 mV, z_q = 0.26±0.02; pH_ex 5.5: V_0.5 = −126.4±16.6 mV, z_q = 0.27±0.04). The fluorescence amplitudes 1-ΔF/F were normalized to the difference between the saturation values at positive or negative potentials, respectively, as obtained from the fits. (D) Reciprocal time constants (τ⁻¹) from fits of a single exponential function to voltage jump-induced fluorescence changes under K⁺-free conditions at pH_ex 5.5 (□) and pH_ex 7.4 (▪). Data are means ± S.E. from 15–17 oocytes. (E,F) Graphs showing the forward (k_f) and reverse (k_b) rate constants of the E₁P↔E₂P transition at pH_ex 7.4 (E) and pH_ex 5.5 (F), as calculated from the observed k_tot = τ⁻¹ in (D) and the voltage-dependent fluorescence amplitudes in (C) according to Supporting Information (Appendix S1). Superimposed in (E,F) are fits of a single exponential function to the calculated k_f and k_b values. The resulting fit parameters (rate constants at 0 mV: k_b(0), k_f(0), and z_q values), as summarized in Table 1, were: pH_ex 7.4 (E): k_f(0) = 2.61±0.05 s⁻¹, z_q,f = −0.06±0.01; k_b(0) = 2.31±0.10 s⁻¹, z_q,b = 0.21±0.01; pH_ex 5.5 (F): k_f(0) = 4.56±0.05 s⁻¹, z_q,f = −0.056±0.003; k_b(0) = 1.22±0.10 s⁻¹, z_q,b = 0.230±0.004.

https://doi.org/10.1371/journal.pone.0033645.g002

Although it would be desirable to study the H⁺ pump under physiological working conditions (i.e. pH_ex down to ∼1), it was not possible to apply such large [H⁺] gradients in our experiments. The recording of a single set of VCF traces as shown in Fig. 2A requires absolutely stable fluorescence for at least 90 s, but at pH values lower than 5.5 the signal quality was too poor for kinetic analyses. Furthermore, as we show below, extracellular acidification leads to significant proton leakage into the cells, which not only impairs long-term stability of the cells, but also leads to an ill-defined [H⁺] gradient. For the sake of reliable working conditions, we had to restrict our analyses to the pH_ex range between 7.4 and 5.5, which - in terms of H⁺ concentration - is still an about 100-fold difference.

A change in the pH_ex from 7.4 to 5.5 altered the fluorescence signals profoundly. At pH_ex 7.4 (Fig. 2A), the largest fluorescence changes occurred at positive voltages, at which the transition to E₂P should be favored, whereas the opposite was observed at pH_ex 5.5 (Fig. 2B), with fluorescence changes being largest upon negative voltage steps, which should drive the enzyme into E₁P. Plotting the steady-state fluorescence amplitudes against the membrane potential resulted in sigmoidal (1-ΔF/F)-V distributions, which could be approximated by a Boltzmann-type function (Fig. 2C). Of note, the pH_ex change from 7.4 to 5.5 resulted in a strong negative V_0.5 shift of the (1-ΔF/F)-V distribution by −105 mV (Fig. 2C and Table 1). This observation is puzzling since it would indicate that an increase of the extracellular proton concentration increases formation of E₂P, in contrast to established paradigms about the electrogenicity of Na,K-ATPase. For the Na⁺ pump, an increase of [Na⁺]_ex shifts the voltage-dependent distribution of the slow charge from ouabain-sensitive transient currents (Rakowski, 1993, Holmgren et al. 2000) towards positive potentials. Hence, high [Na⁺]_ex drives the E₁P/E₂P equilibrium of the Na⁺ pump towards E₁P, in line with the concept of an extracellular access channel for Na⁺ ions.

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Table 1. Parameters characterizing the voltage dependence of the E1P↔E2P conformational transition.

https://doi.org/10.1371/journal.pone.0033645.t001

pH effects on presteady-state kinetics of gastric H,K-ATPase

To delineate the processes underlying this behavior of the H,K-ATPase, we analyzed the kinetics of the voltage step-induced conformational changes. Figure 2D shows the voltage dependence of the reciprocal time constants (τ⁻¹) obtained from fitting single exponential functions to the fluorescence signals at the two different pH_ex values. Assuming that the fluorescence signals directly reflect the redistribution between E₁P and E₂P in response to voltage steps, a simplified two-state kinetic model can be used to derive information about the kinetics of the forward and the backward reaction (see Appendix S1 and [40]). Within this framework, the observed reciprocal time constants (τ⁻¹ = k_tot) represent the sum of the voltage-dependent rate constants for the forward (k_f) and the backward reaction (k_b) of the E₁P↔E₂P transition (τ⁻¹ = k_tot = k_f+k_b), which is coupled to extracellular cation uptake or release steps. From the actual poise of the E₁P/E₂P distribution (reflected by the (1-ΔF/F)-V curve) and k_tot, the individual forward and backward rate constants k_f and k_b at each membrane potential V can be calculated (Eq. A6 and Eq. A7 in Appendix S1). Subsequently, the voltage-dependent values k_f(V) and k_b(V) can be fitted by a single exponential function (according to Eq. A1 and Eq. A2 in Appendix S1). From these fits, the parameters characterizing the voltage dependence of k_f and k_b can be determined, such as the values for the equivalent charge, z_q,f and z_q,b, and the rate constants at 0 mV membrane voltage, k_f(0) and k_b(0). Table 1 summarizes these parameters for all data sets analyzed in this work. Notably, at strongly negative potentials, the reciprocal time constants (τ⁻¹ = k_tot) were similar for both pH_ex values (Fig. 2D). Since the total rate constant k_tot at negative potentials should mainly be determined by k_b of the backward reaction, this observation indicates that extracellular acidification does not accelerate the reverse reaction (E₂P→E₁P). The calculated k_b values (Fig. 2E and 2F) are even lower at acidic pH_ex, with k_b(0) values showing a reduction by about 50% upon a change from pH_ex 7.4 to 5.5 (Table 1). At positive voltages, however, the reciprocal time constants (τ⁻¹ = k_tot) at pH_ex 5.5 were nearly two-fold larger than at pH_ex 7.4 (Fig. 2D), which is reflected by a similar increase in k_f (Fig. 2E,F and Table 1 for k_b(0) and k_f(0) values). These observations indicate an acceleration of the forward reaction (E₁P→E₂P) by extracellular acidification, again contradicting the expectations from an extracellular access channel concept. The slope factors z_q,b, which characterize the voltage dependence of the k_b values in Fig. 2E and 2F, are very similar. Notably, the backward rate constant carries most of the voltage dependence (z_q,b values 0.21 to 0.23, similar to the slope factor z_q from fits of the corresponding (1-ΔF/F)-V curves with a Boltzmann-type function), whereas the forward reaction is only weakly voltage-dependent (z_q,f values 0.05 to 0.06).

Effects of intracellular pH changes on the E₁P/E₂P distribution

Since the H,K-ATPase binds protons intracellularly at around pH 7 but releases them extracellularly against a luminal pH of up to ∼1, it is conceivable that the proton pump might be rather unaffected by pH_ex changes from 7.4 to 5.5, whereas the enzyme may be much more sensitive to minute pH changes at the intracellular side. Therefore, we asked whether the observed V_0.5 shifts of the (1-ΔF/F)-V curves could be due to an influence of the extracellular pH on the pH inside the oocytes. In fact, several studies on Xenopus oocytes reported small pH_in changes upon extracellular acidification [35], [41], [42], with pH_ex 5.5 causing a drop in the intracellular pH by about 0.5 units [42]. To test this hypothesis experimentally, we carried out an “acid-bath procedure” by adding the weak organic acid butyric acid to the extracellular solution. Butyric acid can permeate the plasma membrane in its neutral form and dissociate intracellularly, thereby allowing a controlled intracellular acidification to be achieved (see Material and Methods). To acidify the oocyte interior by ∼0.5 pH units, 40 mM NaCl was replaced by an equal amount of Na-butyrate at pH_ex 7.4, as pioneered elsewhere [35]. In Figure 3, the effects on the voltage-dependent fluorescence signals after a solution exchange from butyrate-free (Fig. 3A) to a butyrate-containing solution (Fig. 3B), and back to butyrate-free solution (Fig. 3C,D) are shown. Notably, the fluorescence changes at pH_ex 7.4 in presence of 40 mM butyrate (Fig. 3B) were very similar to the ones observed at pH_ex 5.5 (Fig. 3E). The (1-ΔF/F)-V curves (Fig. 3F) show that at the same pH_ex of 7.4, the V_0.5 value of the conformational distribution in presence of butyrate is shifted to negative potentials in essentially the same way as observed at pH_ex 5.5 in butyrate-free solution. This supports the hypothesis that the observed V_0.5 shift of the (1-ΔF/F)-V distribution at pH_ex 5.5 entirely results from intracellular acidification. The concept that the H,K-ATPase is tightly regulated by intracellular pH is further supported by the fact that the reciprocal rate constants of the E₁P↔E₂P relaxation in the presence of 40 mM butyrate at pH_ex 7.4 were very similar to those measured at pH_ex 5.5 (Fig. 3G). As already outlined for the rate constants at pH_ex 5.5 (Fig. 2F), the observed increase in k_tot at positive potentials must largely be due to an increased rate constant k_f for the forward reaction, for which a dependence on intracellular pH is rather straightforward. The effects of butyrate on intracellular pH and the conformational distribution of the gastric H,K-ATPase were strictly reversible. Already a few minutes after a solution exchange to butyrate-free solution (Fig. 3C,D), the fluorescence signals were almost identical to the initially observed fluorescence signals at pH_ex 7.4 (Fig. 3A), in agreement with a time constant of ∼315 s determined by Stewart et al. for the recovery of intracellular pH after withdrawal of butyrate [35].

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Figure 3. Effects of intracellular acidification on the E₁P/E₂P distribution and Rb⁺ transport.

(A–E) Fluorescence responses of TMRM-labeled HKαS806C/βWT under extracellular K⁺-free conditions (90 mM extracellular Na⁺) upon voltage jumps from −40 mV to potentials between −180 mV and +60 mV in −20 mV steps. Recordings in (A–D) originated from a single oocyte (A) at pH_ex 7.4, (B) after 1 min in presence of 40 mM Na-butyrate (pH_ex 7.4), and after 1 min (C) and 2 min (D) washout of butyrate (pH_ex 7.4 buffer). (E) Fluorescence responses from a different cell in pH_ex 5.5 buffer. (F) Voltage dependence of stationary fluorescence amplitudes 1-ΔF/F from the recordings in (A–E) at pH_ex 7.4 (▪), pH_ex 7.4+40 mM butyrate (□), and after 1 min (⋄) or 2 min (▵) washout of butyrate. Data at pH_ex 5.5 (•) are also shown. Fits of a Boltzmann-type function are superimposed to each data set, and the fluorescence amplitudes were normalized to saturation values from the fits. (G) Reciprocal time constants (τ⁻¹) from fits of a single exponential function to fluorescence changes under K⁺-free conditions. Data obtained at pH_ex 7.4 in the presence of 40 mM butyrate (♦) are compared to those in butyrate-free solutions at pH_ex 5.5 (□) and at pH_ex 7.4 (▪). Data are means±S.E. from 12–17 oocytes. (H) H,K-ATPase-mediated Rb⁺ uptake (at 5 mM Rb⁺) measured on individual cells by atomic absorption spectroscopy in the absence (gray) or presence (black) of 10 µM SCH28080 at different pH_ex and ionic conditions. Results from non-injected and HKαS806C/βWT-expressing oocytes at pH_ex 5.5, pH_ex 7.4, and pH_ex 7.4+40 mM butyrate are shown. Data are means±S.E. from three experiments on different cell batches with 15–20 oocytes per condition, and normalized to the Rb⁺ uptake of HKαS806C/βWT at pH_ex 5.5 (mean specific activities of 15.4, 23.1 and 39.0 pmol/oocyte/min).

https://doi.org/10.1371/journal.pone.0033645.g003

Effect of extra- and intracellular pH on steady-state cation pumping

To test the significance of the described pH effects on stationary H⁺/K⁺ exchange transport, we measured the Rb⁺ transport activity of the H,K-ATPase under turnover conditions at saturating Rb⁺ concentrations (5 mM). As shown in Figure 3H, Rb⁺ uptake of the gastric H,K-ATPase at pH_ex 5.5 was more than two-fold larger than the transport activity at pH_ex 7.4. Again, the effect of the extracellular acidification could be attributed to an intracellular pH decrease, since almost the same increase in transport activity was observed at pH_ex 7.4 in presence of 40 mM butyrate. These findings strongly suggests that the availability of protons at the intracellular side is not only rate-limiting for the E₁P→E₂P transition (Fig. 3G), but also for the turnover rate during stationary cation pumping. Notably, the relatively increased E₁P preference of the enzyme at neutral intracellular pH (pH_ex 7.4 without butyrate, Fig. 2C) compared to conditions of slight intracellular acidification (pH_ex 5.5 or pH_ex 7.4 with butyrate) was also reflected by distinct differences regarding the inhibition by 10 µM SCH28080 (black bars in Fig. 3H). To understand the effects of this reversible, E₂/E₂P-specific, K⁺-competitive inhibitor, two aspects have to be kept in mind. First it must be considered that only the protonated form of SCH28080 is pharmacologically active, with deprotonation occuring with a pK_a of 5.5 [43], [44]. Consequently, at pH_ex 7.4 only about 1% of the total inhibitor concentration (i.e. ∼0.1 µM) is in the protonated, active form, which is in the same range as the IC₅₀ values determined for this K⁺-competitive antagonist (67 nM for [K⁺] = 0, and 480 nM for [K⁺] = 10 mM [45]; 0.18 µM and 0.66 µM for K⁺-stimulated ATPase activity [43]), whereas at pH_ex 5.5 about 50% of the compound is active. The second important point is the fractional amount of enzyme molecules in E₂ or E₂P, because the inhibitor is specific for these intermediates. Therefore, the highly effective inhibition at pH_ex 5.5 is on one hand due to the higher abundance of the active compound, and on the other hand due to the strong shift towards the E₂P-state (at potentials around −10 mV, which are relevant for the Rb⁺ flux measurements without voltage control, see Fig. 2C and 3F) caused by the concomitant decrease in intracellular pH. However, the higher extent of SCH28080 inhibition at the same pH_ex of 7.4, depending on whether the cytoplasm is acidified by the addition of butyrate or not (Fig. 3H, black bars), indicates that intracellular acidification indeed entails a higher E₂P preference of the H,K-ATPase.

Changes in the conformational distribution in response to extracellular K⁺

Of note, all VCF experiments shown so far were done under K⁺/Rb⁺-free conditions and therefore do not reflect the conditions of the Rb⁺ fluxes in Fig. 3H. In order to discriminate the effects of different monovalent cations on the E₁P/E₂P distribution, we first measured the changes of the voltage-dependent fluorescence signals under H⁺/K⁺ turnover conditions (Fig. 4). After a change from K⁺-free solution to 5 mM K⁺, the magnitudes of fluorescence changes were substantially reduced at both investigated pH_ex values (Fig. 4A,B), with the effect being more pronounced at pH_ex 7.4 (compare black bars in Fig. 4D). This is very similar to the effect of K⁺ on the fluorescence changes of TMRM-labeled Na,K-ATPase [31]. Cyclic turnover at high K⁺ concentrations results in a redistribution of enzyme molecules over all reaction cycle intermediates, thereby increasing the accumulation of states (e.g. dephosphorylated E₁-type intermediates), whose occupancies are insensitive to transmembrane voltage. This, in effect, diminishes the number of pump molecules that contribute to the fluorescence changes related to the voltage-dependent redistribution between E₁P and E₂P states. Due to the proposed acceleration of the rate-limiting step early within the H⁺ branch of the cycle by slight intracellular acidification (as a result of the pH_ex change to 5.5), fewer molecules are kinetically trapped in voltage-insensitive intermediates, which explains the significantly larger fluorescence changes under turnover conditions at pH_ex 5.5. A comparison of the time course of the fluorescence responses to −180 mV and +60 mV from Figure 4A in the absence of K⁺ and at 5 mM K⁺ (see normalized signals in Fig. 4C) shows that K⁺ accelerates the conformational relaxation at negative as well as positive potentials. This global acceleration of rate constants indicates that K⁺ opens up a second relaxation pathway (E₂P→E₂→E₁) that occurs in addition to the E₁P↔E₂P relaxation.

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Figure 4. Effects of extracellular K⁺ on voltage-dependent fluorescence changes.

(A,B) Voltage step-induced fluorescence responses of TMRM-labeled oocytes expressing HKαS806C/βWT in K⁺-free (upper traces) or 5 mM K⁺-containing extracellular solution (lower traces) at pH_ex 5.5 (A), and pH_ex 7.4 (B), according to a voltage protocol as in Fig. 2A (inset). (C) Comparison of the time course of the fluorescence signals from panel (A) in response to voltage pulses to −180 mV and +60 mV in the absence of K⁺ and in the presence of 5 mM K⁺. Signals were normalized to the fluorescence amplitude reached at the end of the voltage pulse to −180 mV or +60 mV, respectively. (D) Comparison of normalized ΔF/F values (change in stationary fluorescence between −180 mV and +60 mV, divided by fluorescence at −40 mV) in the absence (gray bars) or presence (black bars) of 5 mM extracellular K⁺, both at pH_ex 5.5 and pH_ex 7.4. Data are means±S.D. from 3–5 oocytes, normalized to the mean ΔF/F at pH_ex 5.5 in K⁺-free solution.

https://doi.org/10.1371/journal.pone.0033645.g004

Na⁺ effects on presteady-state fluorescence changes of gastric H,K-ATPase

In a previous publication, we found indications for a competition between Na⁺ and Rb⁺ at the extracellular binding sites [28], since the apparent affinity for extracellular Rb⁺ in Rb⁺ uptake experiments was reduced about 7-fold in the presence of extracellular Na⁺. To scrutinize, whether such competitive effects of extracellular Na⁺ ions also affect the voltage dependence and kinetics of the E₁P↔E₂P transition, we compared the voltage dependence of the fluorescence amplitudes and of the respective reciprocal time constants in extracellular Na⁺-free and Na⁺-containing solutions (Fig. 5). Notably, the parameters z_q for the (1-ΔF/F)-V distributions were larger in the absence than in the presence of Na⁺ at both pH_ex values (Fig. 5A,B), which will be rationalized in the Discussion and Supporting Information (Appendix S2 and Appendix S3). Furthermore, the presence of extracellular Na⁺ had a large effect on the V_0.5 values of the E₁P/E₂P distribution at pH_ex 7.4 (Fig. 5A and Table 1), but not at pH_ex 5.5 (Fig. 5B and Table 1). From the voltage dependence of the reciprocal time constants (τ⁻¹ = k_tot) (Fig. 5C,F) and the k_f and k_b values calculated thereof (Fig. 5D,E and Fig. 5G,H), it is evident that extracellular Na⁺ accelerates the reverse rate constants k_b(0) by a factor of ∼2.5 at both pH_ex values. This indicates that extracellular Na⁺ ions, which are by far more abundant than protons at pH_ex 7.4 as well as 5.5, can act as H⁺ analogs when the binding sites face the extracellular medium (see Discussion). In contrast, k_f(0) is only slightly changed (∼22% decrease at pH_ex 7.4, Fig. 5D,E; ∼15% increase at pH_ex 5.5, Fig. 5G,H) compared to Na⁺-free conditions (Table 1). Thus, extracellular Na⁺ mainly accelerates the reverse rate constant k_b, whereas k_f is essentially unchanged, and the total rate constant is consequently increased only at negative voltages, in notable contrast to the effect of K⁺ on the relaxation kinetics (see Discussion).

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Figure 5. Effect of extracellular Na⁺ on the E₁P↔E₂P conformational transition.

(A,B) Voltage dependence of fluorescence amplitudes 1-ΔF/F of TMRM-labeled HKαS806C/βWT at pH_ex 7.4 (A), and pH_ex 5.5 (B) in the presence of 90 mM extracellular Na⁺ (▪,□) compared to Na⁺-free conditions (•,○; Na⁺ replacement by 90 mM TMA⁺). Data are means±S.E. of 13–15 oocytes. Superimposed are curves resulting from a fits of a Boltzmann-type function to the data (pH_ex 7.4, 90 mM TMA⁺: V_0.5 = −89.6±3.3 mV, z_q = 0.48±0.04; pH_ex 7.4, 90 mM Na⁺: V_0.5 = −19.7±5.4 mV, z_q = 0.26±0.02; pH_ex 5.5, 90 mM TMA⁺: V_0.5 = −125.2±11.4 mV, z_q = 0.49±0.07; pH_ex 5.5, 90 mM Na⁺: V_0.5 = −126.4±16.6 mV, z_q = 0.26±0.03), parameters are listed in Table 1. The fluorescence amplitudes were normalized to the saturation values from the fits. (C,F) Reciprocal time constants (τ⁻¹) from fits of a single exponential function to fluorescence signals in Na⁺-free and in 90 mM Na⁺-containing solutions for pH_ex 7.4 (C) and pH_ex 5.5 (F). Data are means±S.E. from 13–15 oocytes. (D,E) Calculated forward (k_f) and reverse (k_b) rate constants of the E₁P↔E₂P transition in the presence of 90 mM Na⁺ (D), and in Na⁺-free solution (E) at pH_ex 7.4, as calculated from the observed k_tot = τ⁻¹ values in (C) and the voltage-dependent fluorescence amplitudes in (A) according to Supporting Information (Appendix S1). (G,H) Calculated forward (k_f) and reverse (k_b) rate constants at pH_ex 5.5 in the presence of 90 mM Na⁺ (G) and in Na⁺-free solution (H), as calculated from the k_tot = τ⁻¹ values in (F) and fluorescence amplitudes in (B). Superimposed in (D,E,G,H) are fits of a single exponential function to the k_f and k_b values, the resulting fit parameters (k_b(0), k_f(0), and z_q values) are summarized in Table 1.

https://doi.org/10.1371/journal.pone.0033645.g005

Temperature dependence of steady-state pump activity

Rb⁺ uptake measurements were carried out at different temperatures to determine the activation energy of Rb⁺ transport at pH_ex 5.5 and pH_ex 7.4. As shown in Figure 6A, the Rb⁺ transport activity at pH_ex 5.5 was substantially larger than at pH_ex 7.4 in the whole temperature range covered by our experiments (18–34°C). Arrhenius plots yielded linear relationships at both investigated pH (Fig. 6B). At pH_ex 7.4, we consistently observed in several independent experiments that the data points corresponding to a temperature of 34°C significantly diverged from the linear function defined by the other data points. Such a behavior is not uncommon, as exemplified by the temperature dependence of K⁺-stimulated pump currents of Na,K-ATPase expressed in oocytes, for which a reduced slope of the Arrhenius plot at temperatures above 26°C was observed too [46]. Exclusion of the data point for 34°C at pH_ex 7.4 yielded activation energies of similar magnitude at both pH_ex conditions (95.8±1.7 kJ at pH_ex 5.5 versus 91.7±3.7 kJ at pH_ex 7.4). The close similarity of these values suggests that Rb⁺ uptake of the gastric proton pump at both pH values is rate-limited by the same partial reaction, and due to the high activation energy, this step is likely not to be diffusion-controlled, but might be related to a major conformational change.

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Figure 6. Temperature and voltage dependence of Rb⁺ uptake by gastric H,K-ATPase.

(A) H,K-ATPase-mediated Rb⁺ uptake (in pmol/oocyte/min) at 5 mM Rb⁺ and a pH_ex of 7.4 (light gray bars) or 5.5 (gray bars) at temperatures between 18 and 34°C, as indicated. White bars represent Rb⁺ uptake of non-injected control oocytes at each temperature and pH_ex 5.5. The black bar at 34°C shows the residual Rb⁺ uptake at pH_ex 5.5 in the presence of 100 µM SCH28080. Data in each column are means of 20–25 oocytes from oocytes of one cell batch. (B) Arrhenius plot for temperature-dependent Rb⁺ uptakes from data as in (A) at pH_ex 7.4 (▪), and pH_ex 5.5 (○). Data represent means±S.E. of three independent experiments (similar to the one shown in A), after normalization to Rb⁺ uptake at 34°C for each experiment. Activation energies obtained from linear fits to the data (superimposed lines) are given for each pH_ex. (C) Rb⁺ uptake (in pmol/oocyte/min) at 5 mM Rb⁺ and pH_ex 7.4 or 5.5 for oocytes expressing HKαS806C/βWT, which had either been clamped to a membrane potential of −100 mV, or subjected to Rb⁺ uptake without voltage clamping (V_m∼−10 to −20 mV). Black bars represent Rb⁺ uptake of H,K-ATPase-expressing oocytes clamped at −100 mV in the presence of 100 µM SCH28080. Data are means±S.D. from several oocytes of a single batch (numbers stated on each column).

https://doi.org/10.1371/journal.pone.0033645.g006

Voltage dependence of steady-state cation transport

To assess the voltage dependence of the overall pump activity, we performed Rb⁺ uptake experiments also under transmembrane voltage control (Na⁺-free conditions). In Figure 6C, the Rb⁺ uptake activity at saturating Rb⁺ concentrations (5 mM) in not voltage-clamped oocytes (V_m∼−10 to −20 mV, determined in independent experiments) is compared to the Rb⁺ uptake activity of oocytes whose membrane potential was clamped to −100 mV by two-electrode voltage clamping. At pH_ex 7.4 as well as pH_ex 5.5, only a slight and hardly significant decrease of the Rb⁺ transport activity was observed at −100 mV compared to unclamped oocytes. Importantly, however, the about two-fold increase of Rb⁺ transport at pH_ex 5.5 compared to pH_ex 7.4, as observed previously in Fig. 3H, occurred irrespective of the membrane potential. This finding supports the hypothesis that an intracellular pH-sensitive and only weakly voltage-dependent event is not only rate-limiting for the E₁P→E₂P conformational transition (monitored by the VCF experiments) but also for the overall pumping rate.

Effects of extracellular pH on the E₁P/E₂P distribution

It is generally accepted that the transport cycle of the H,K-ATPase proceeds according to a Post-Albers-type reaction scheme, as formulated for the Na,K-ATPase, despite some difference in detail, such as the strong E₂P preference of the gastric proton pump under physiological conditions [26], [28], [47], [48]. Although the H,K-ATPase carries out net electroneutral transport, experiments using H,K-ATPase-containing parietal cell membrane fragments attached to black lipid membranes have shown transient current signals upon ATP concentrations jumps in the absence of K⁺ [19], [20], [21] suggesting that an electrogenic event takes place during H⁺ translocation. Thus, as a first approach, one could assume that electrogenicity in the H,K- and the Na,K-ATPase follows the same mechanism. For the Na⁺ pump, the slowest phase of presteady-state Na⁺ movement, which is kinetically coupled to the E₁P↔E₂P transition, arises from extracellular Na⁺ release from (or reverse binding to) a site located at ∼70% of the electrical distance from the extracellular side. According to the high-field access channel hypothesis, changes in membrane potential are kinetically equivalent to changes in the ‘effective’ ion concentration at the binding sites deep within the ion well [49], [50]. Thus, an about 100-fold increase in the extracellular H⁺ concentration (change from pH_ex 7.4 to 5.5) should shift the V_0.5 value of the E₁P/E₂P distribution towards E₁P. The resulting shift (ΔV_0.5) could then be predicted from a Nernst-like equation [9], [12], [50]:(1)Thus, using an equivalent charge or fractional well depth (z_q) of 0.26–0.27 as derived from the Boltzmann curve parameters in Fig. 2C, a pH_ex change from 7.4 to 5.5 should result in a positive ΔV_0.5 of 415 mV. However, in contrast to the expectations for an extracellular H⁺ access channel, our VCF data show that a ΔpH_ex of 1.9 units shifts V_0.5 by about −105 mV.

To understand why a ΔpH_ex of 1.9 units does not cause a shift towards E₁P, it must be considered that the H,K-ATPase in situ releases protons against a luminal pH below 1 ([H⁺]∼150 mM in the stomach), which implies that extracellular proton release from the binding pocket occurs with a pK_a value even lower than 1. Thus, even at a pH_ex of 5.5, the proton concentration is by several orders of magnitude too small to achieve a sufficient occupancy at the luminal-facing H⁺-binding site(s), which would be a prerequisite for an E₁P shift of the E₁P/E₂P distribution. Even a transmembrane voltage of −200 mV would increase the effective proton concentration at the bottom of an extracellular access channel with a fractional depth z_q of 0.26 (Fig. 2C) by only about 8-fold, resulting in a still insufficient ‘effective’ pH of 4.6, which is still far from the physiological pH of ∼1–2. With Na,K-ATPase, the conditions for the study of electrogenic Na⁺ transport are more favorable, since. the extracellular Na⁺ affinity of the Na⁺ pump is in the order of several hundreds of mM [7], [12], and Na⁺ concentrations in this range can easily be applied in electrophysiological experiments. Unfortunately, pH_ex 1 (equivalent to [H⁺] = 100 mM) cannot be tested in Xenopus oocyte experiments so that the question whether protons traverse an extracellular ion well cannot be resolved. However, due to the effects of extracellular Na⁺ ions on the conformational distribution the existence of an extracellular access channel of the H,K-ATPase cannot be ruled out, as discussed below.

Intracellular pH strongly influences kinetics and poise of the E₁P/E₂P distribution

To explain the negative V_0.5 shift of the (1-ΔF/F)-V distribution in response to pH changes (Fig. 2C), our experiments designed to achieve a controlled intracellular acidification show that the observed V_0.5 shift can entirely be attributed to a slight intracellular acidification that is induced by an extracellular pH change. In fact, the (1-ΔF/F)-V curves (Fig. 3F) as well as the reciprocal rate constants (Fig. 3G) of the fluorescence signals at pH_ex 5.5 and at pH_ex 7.4 in the presence of 40 mM butyrate (which lowers pH_in by ∼0.5 units) are fully equivalent. Therefore, the extracellular pH (between 7.4 and 5.5) is apparently irrelevant for the poise of the E₁P/E₂P distribution, whereas already a small deviation from a neutral intracellular pH produces a large effect. The dependence on pH_in is in line with an intracellular access channel for protons. Indeed, the calculation of ΔV_0.5 (Eq. 1) according to an intracellular pH change by 0.4–0.5 units and an ‘effective’ z_q of 0.26–0.27 (in the presence of 90 mM extracellular [Na⁺]) yields a ΔV_0.5 of about −90 to −110 mV, which is in good agreement to the observed shift of −105 mV (Fig. 2C and 3F). For the (1-ΔF/F)-V distributions measured in the absence of extracellular Na⁺ (Fig. 5A,B), the observed ΔV_0.5 (−35 mV) agrees less well with theoretical values (−48 to −61 mV, with z_q between 0.48 and 0.49). However, considering the strongly negative V_0.5 values of the (1-ΔF/F)-V curves in question, it must be noted that oocyte TEVC experiments at voltages below −180 mV become increasingly problematic.

Extracellular Na⁺ ions compete with protons for access to E₂P, and kinetic analysis elicits the fractional depth of intra- and extracellular access channels

Since extracellular Na⁺ ions not only reduce the apparent affinity for extracellular Rb⁺ as K⁺ congeners in Rb⁺ uptake studies [28], but also profoundly change the conformational distribution (compared to the relatively small effect achieved by extracellular acidification, Fig. 5A,B), an extracellular cation access channel still has to be considered for the H,K-ATPase. At pH_ex 7.4, a Na⁺ concentration of 90 mM leads to a stronger accumulation of E₁P at physiological potentials (around −70 mV), as actually expected for high [H⁺], by a combined effect on V_0.5 and a decrease in the slope factor z_q (Fig. 5A,B). The larger fraction of E₁P correlates with an increase of the reciprocal rate constants at hyperpolarizing potentials (Fig. 5C), which indicates an increase of the rate constant for the backward reaction (Fig. 5D,E), whereas the forward rate constant is hardly changed. These observations agree with the notion that Na⁺ ions, which are 10⁴- to 10⁶-fold more abundant than protons in our experiments, can act as H⁺ analogs within an extracellular-facing ion well. At pH_ex 5.5, the Na⁺ effect on V_0.5 of the conformational distribution was no longer present (Fig. 5B), although the reciprocal time constants at negative potentials were also increased (Fig. 5F) suggesting that the effect of Na⁺ ions on the E₁P↔E₂P kinetics is present even upon a 100-fold increase of the extracellular [H⁺]. But, at this lower pH_ex of 5.5, the E₁P-shifting effect of the increased k_b values is counteracted by the simultaneous increase of the forward rate constants (Fig. 5G,H) that occurs due to the intracellular acidification. The fact that Na⁺ ions exert H⁺-like effects on H,K-ATPase is another example for the promiscuity of the external-facing cation binding sites in P-type pumps, as outlined recently for Na,K-ATPase, in which some alkali metal ions or monovalent organic cations were shown to induce Na⁺-like or K⁺-like functional effects [51].

Since it is reported in the literature that for both Na,K- and H,K-ATPase Na⁺ ions can mimic the effect of K⁺ ions in the dephosphorylation limb of the cycle [27], one could argue that the observed kinetic effects of Na⁺ on the conformational distribution might be due to an alternative reaction branch. However, if Na⁺ ions would act like K⁺ ions to stimulate the E₂P→E₂→E₁ pathway, Na⁺ addition should result in a global increase of the total relaxation rate constant (as indeed seen for K⁺, Fig. 4C), which is not observed. In fact, Na⁺ mainly affects k_b, but not k_f, and thus increases the total rate only at negative potentials (Fig. 5). Furthermore, a significant entry of enzyme molecules into the K⁺ limb of the cycle should lead to an accumulation of E₁ states, which cannot contribute to the voltage-dependent E₁P↔E₂P relaxation. Thus, similar to the results of K⁺ addition in Fig. 4A,B, the absolute fluorescence amplitudes should decrease, which is also not observed with Na⁺. Therefore, we conclude here that under the conditions of our experiments there is no indication for a significant effect of Na⁺ on the dephosphorylation branch of the H,K-ATPase cycle.

Notably, at both investigated pH_ex, Na⁺ had a strong effect on the slope factor z_q of the (1-ΔF/F)-V distribution (∼0.26–0.27 with, versus ∼0.48–0.49 without extracellular Na⁺ ions). As outlined in Supporting Information (see Appendix S2, Appendix S3, and Figure S1), such a situation can arise from a superposition of effects resulting from cation binding through an intra- and an extracellular access channel. For the pseudo three-state model depicted in Fig. 1B, the following assumptions are made: First, the z_q factor of ∼0.5 measured in the absence of external Na⁺ exclusively represents the fractional depth of an intracellular H⁺ access channel. Second, external Na⁺ ions exert their effect on the conformational distribution by binding through a shallower extracellular ion well with a z_q of ∼0.2. This is reasonable, since the H,K-ATPase lacks the third ‘unique’ cation binding site characteristic for the Na⁺ pump, which is responsible for the major electrogenic release of the third Na⁺ ion with a fractional charge of ∼0.8, whereas the release/uptake of cations to the two ‘common’ sites occurs with a smaller apparent valence of ∼0.2. The model simulations in Appendix S3 qualitatively reproduce the experimental observations (Figure S1): First, the inclusion of an additional electrogenic extracellular Na⁺ uptake step enforces a positive shift in V_0.5. Second, such uneven z_q factors distort the voltage dependence of the resultant conformational distribution in a way that fitting by a simple Boltzmann-type function yields an ‘effective’ z_q value of even less than 0.5 (Appendix S3 and Figure S1), exactly as observed in Fig. 5A.

Thus, the voltage dependence of the calculated rate constants k_f and k_b of the forward and backward reaction (Fig. 2E,F and Fig. 5D,E,G,H) can be reconciled with the concept of a high-field intracellular and a shallower extracellular access channel. Although more detailed kinetic information would be required to correlate the calculated rate constants k_f and k_b with individual rate constants within the pseudo three-state model of Appendix S2, a tentative assignment appears feasible. The data in Fig. 5D,E,G,H show that one of the rates, k_b, is rather strongly dependent on membrane potential (with z_q values of ∼0.23), whereas the voltage dependence of the other, k_f, is very weak. In case of the Na⁺ pump, the voltage insensitivity of the forward rate constant from ouabain-sensitive transient currents is attributed to a voltage-independent reaction step (the E₁P→E₂P conformational transition in conjunction with Na⁺ deocclusion) that is rate-limiting the subsequent Na⁺ release step(s). The increase of rate constants upon hyperpolarization results from the fact that negative potentials favor the entry of Na⁺ ions to the binding pocket through an extracellular high-field access channel. With an hypothetical intracellular access channel in the case of H,K-ATPase, the fact that the forward rate constant k_f is independent from voltage (and pH_ex) suggests that a voltage-independent step preceding intracellular H⁺ binding (step 1 in Fig. 1A) is rate-limiting for H⁺ uptake and the subsequent E₁P→E₂P conformational transition. Conversely, the relatively steep voltage dependence of k_b results from the fact that negative voltages speed up the intracellular release of H⁺ through the access channel. The [Na⁺]_ex-dependence of the partial reaction represented by k_b might be the consequence of Na⁺ ions traversing a shallow extracellular access channel to reach the binding pocket, which, in effect, will also speed up the E₂P→E₁P transition.

Proposed mechanism for the effect of intracellular pH on the H⁺/K⁺ pumping rate

The dependence of the proposed voltage-independent step preceding intracellular H⁺ binding on the intracellular pH could mean that prior to the electrogenic binding of H⁺ to the transport site(s) a proton must bind to a ‘regulatory’ site (with a pK_a around neutral), which is in rapid equilibrium with the intracellular pH. Neutralization of a protonatable residue might change the local electrostatics, eventually leading to the formation of the access channel itself or to control the accessibility of the ion well for intracellular protons. Alternatively, the ‘regulatory’ proton could even be one of the presumably two protons that are transported in each reaction cycle. Of note, our Rb⁺ uptake experiments suggest that the availability of protons at the intracellular side is not only rate-limiting for the E₁P→E₂P conformational transition (Fig. 3G), but also for the turnover rate during stationary cation pumping (Fig. 3H). This conclusion can be drawn from the fact that both the stationary turnover number (Rb⁺ uptake) and the forward rate constant of the E₁P↔E₂P relaxation (monitored by the VCF experiments) show an about two-fold increase upon intracellular acidification by 0.5 pH units. Importantly, at pH_ex 7.4 as well as pH_ex 5.5, only a very small decrease of the Rb⁺ transport activity was observed at −100 mV compared to unclamped oocytes, whereas the about two-fold increase of Rb⁺ transport at pH_ex 5.5 compared to pH_ex 7.4 occurred irrespective of the membrane potential. The weak voltage sensitivity of cation transport is, first, reflected by the hardly voltage-sensitive rate constants k_f (Figure 5D,E,G,H). Second, if the rate constant for K⁺(Rb⁺)-dependent dephosphorylation is much faster than the voltage-dependent relaxation between E₁P and E₂P, the majority of H,K-ATPase molecules on average will dwell in states (e.g. dephosphorylated intermediates like E₁), whose occupancies are insensitive to transmembrane voltage, as indicated from the small voltage-dependent fluorescence changes under turnover conditions (Fig. 4).

The close similarity of the activation energies at pH_ex 7.4 and 5.5 also suggests that Rb⁺ uptake of the gastric proton pump is rate-limited by the same pH_in-dependent partial reaction, and the high E_A values suggest that this step is likely not diffusion-controlled, but might be related to a major conformational change. The activation energies at pH_ex 7.4 and pH_ex 5.5 reported here are remarkably close to the 93 kJ/mol determined by Stengelin and co-workers in BLM experiments at an intermediate pH of 6.2. [21]. This value was obtained from the temperature-dependence of a time constant (τ₃) that was assigned to the phosphorylation reaction and covered an even larger temperature range between 3°C and 40°C. The agreement between these values corroborates the idea that the overall pump activity monitored by the Rb⁺ uptake measurements is indeed rate-limited by partial reactions of the H⁺ outward moving branch, i.e. the phosphorylation reaction (that strongly depends on the intracellular H⁺ concentration) and the subsequent E₁P→E₂P conformational transition reflected by the presteady-state fluorescence measurements.

Proposed role of Glu-820 for intra- and extracellular proton sensitivity

In a recent study, we have identified an acidic residue belonging to the putative cation binding pocket of the gastric H,K-ATPase (Glu-820 in M6) that might be crucial for the sensitivity towards intracellular acidification described here. Upon replacement of Glu-820 by non-protonatable residues (e.g. Gln or Ala), Rb⁺ uptake did not increase in presence of butyrate at pH_ex 7.4 (see Fig. 6F in [40]), which is very different from the aforementioned behavior of the wild-type enzyme. At pH_ex 5.5, Rb⁺ uptake by the two charge-neutralizing mutants was even reduced indicating that the mutations result in an increased competition of extracellular protons with Rb⁺ ions at the binding sites. Therefore, Glu-820 could be crucial for determining K⁺ (Rb⁺) selectivity in the E₂P state, which is especially important at steep H⁺ gradients. Glu-820 may represent a site were protons are transiently bound before being expelled to the extracellular space, since the proximity of Glu-820 to the charged side-chain of Lys-791 (see Figure S2) could facilitate the large pK changes that are required to enable expulsion of a proton from this site at physiological pH of ∼1 into the stomach lumen. If the site is not occupied by a proton, the two oppositely charged residues Lys-791 and Glu-820 are probably forming a salt bridge that stabilizes the pump in the E₂P state, as proposed earlier [40], [52].

Concluding remarks

In the absence of extracellular K⁺, extracellular acidification from pH_ex 7.4 to 5.5 has no effect on the E₁P↔E₂P relaxation of gastric H,K-ATPase. In contrast, intracellular acidification by ∼0.5 pH units speeds up the forward relaxation rate and increases the H⁺/K⁺ pumping rate two-fold. Extracellular Na⁺ ions compete with protons and K⁺ ions for entry into the extracellular-facing access channel to the binding sites in E₂P, but have no significant effect on the dephosphorylation branch of the cycle. Kinetic analysis based on a pseudo-three state model that simultaneously includes voltage-dependent (un)binding/(de)occlusion steps through an intra- and an extracellular access channel indicates that the intracellular access channel for protons has a fractional depth of ∼0.5, whereas the extracellular access channel, which is accessible for protons, Na⁺ and K⁺ ions, has a fractional depth of ∼0.2. The overall H⁺/K⁺ pumping rate is essentially voltage-insensitive indicating that a voltage-independent step is rate-limiting for the pump cycle. This intracellular pH-sensitive, rate-limiting step might be the intracellular binding of a proton to a regulatory binding site, which could be the transport site, to which the side chain of E820 is contributing.