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¹ Department of Anesthesiology, Pharmacology, and Therapeutics and ² Department of Cellular and Physiological Sciences, University of British Columbia, Vancouver, V6T 1Z3 Canada

Correspondence to David Fedida: fedida{at}interchange.ubc.ca

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Lowering external pH reduces peak current and enhances current decay in Kv and Shaker-IR channels. Using voltage-clamp fluorimetry we directly determined the fate of Shaker-IR channels at low pH by measuring fluorescence emission from tetramethylrhodamine-5-maleimide attached to substituted cysteine residues in the voltage sensor domain (M356C to R362C) or S5-P linker (S424C). One aspect of the distal S3-S4 linker -helix (A359C and R362C) reported a pH-induced acceleration of the slow phase of fluorescence quenching that represents P/C-type inactivation, but neither site reported a change in the total charge movement at low pH. Shaker S424C fluorescence demonstrated slow unquenching that also reflects channel inactivation and this too was accelerated at low pH. In addition, however, acidic pH caused a reversible loss of the fluorescence signal (pKa = 5.1) that paralleled the reduction of peak current amplitude (pKa = 5.2). Protons decreased single channel open probability, suggesting that the loss of fluorescence at low pH reflects a decreased channel availability that is responsible for the reduced macroscopic conductance. Inhibition of inactivation in Shaker S424C (by raising external K⁺ or the mutation T449V) prevented fluorescence loss at low pH, and the fluorescence report from closed Shaker ILT S424C channels implied that protons stabilized a W434F-like inactivated state. Furthermore, acidic pH changed the fluorescence amplitude (pKa = 5.9) in channels held continuously at –80 mV. This suggests that low pH stabilizes closed-inactivated states. Thus, fluorescence experiments suggest the major mechanism of pH-induced peak current reduction is inactivation of channels from closed states from which they can activate, but not open; this occurs in addition to acceleration of P/C-type inactivation from the open state.

	INTRODUCTION

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The kinetic properties of many voltage gated potassium (Kv) channels are affected by external pH. Low extracellular pH reduces the peak current in Shaker channels as well as in Kv1.3, Kv1.4, and Kv1.5 from the related Kv1 family of mammalian channels (Deutsch and Lee, 1989; Lopez-Barneo et al., 1993; Perez-Cornejo, 1999; Steidl and Yool, 1999; Claydon et al., 2000; Jäger and Grissmer, 2001; Kehl et al., 2002; Starkus et al., 2003; Somodi et al., 2004) in a manner that is independent of the depolarizing shift of the voltage dependence of channel opening that is caused by a proton-induced charge screening effect (Deutsch and Lee, 1989; Kehl et al., 2002; Trapani and Korn, 2003). In pH-sensitive mammalian Kv1 channels, a histidine residue in the outer pore is responsible for the inhibition of current and consistent with this, the pK_a (with physiological external K⁺ concentrations) is pH 6.1–6.2 (Steidl and Yool, 1999; Claydon et al., 2000; Kehl et al., 2002). In the case of the Shaker channel, which does not possess a histidine residue within the outer pore, the pK_a is pH 4.7–5.0 suggesting that the pH sensor may be an aspartate or glutamate residue (Perez-Cornejo, 1999; Starkus et al., 2003).

In the absence of N-type inactivation, acidic pH not only reduces peak current amplitude, but also enhances the decay of current in Shaker, Kv1.4, and Kv1.5 channels during the depolarizing pulse (Perez-Cornejo, 1999; Claydon et al., 2002; Starkus et al., 2003; Zhang et al., 2003). It has been suggested that the decrease in peak current observed with extracellular acidification occurs as a direct consequence of the pH-induced enhancement of open-state inactivation, which results in an accumulation of channels in the P/C-type inactivated state (Steidl and Yool, 1999; Starkus et al., 2003; Zhang et al., 2003). In support of this, the acceleration of current decay and the decrease of peak current amplitude at low pH is abolished in Shaker, Kv1.4, and Kv1.5 mutant channels in which inactivation is compromised by mutation of the outer pore residue equivalent to T449 in Shaker (K532 and R487 in Kv1.4 and Kv1.5, respectively), to a valine or tyrosine residue (Claydon et al., 2002; Starkus et al., 2003; Zhang et al., 2003; Kurata and Fedida, 2006).

However, an additional mechanism to explain the decreased peak Kv channel current at low pH has also been proposed from our previous work. During extracellular acidification (to pH 5.9), Kv1.5 current inhibition did not demonstrate use dependence as would be expected if the current was being reduced by an accumulation of open-state inactivation (Kehl et al., 2002). In addition, a 2-min rest interval with pH 5.9 bath solution, which would allow enough time for recovery from pH-induced open-state inactivation, did not relieve the current inhibition, which suggests that inactivated channels cannot recover easily at low pH. This idea of stabilization of closed-state inactivated channels at low pH is supported by a recent study investigating the effect of acidic pH on single channel Kv1.5 current. Acidic pH did not alter the Kv1.5 single channel conductance, but increased the number of null sweeps and induced gating behavior in which the channel switched between available and unavailable modes (Kwan et al., 2006). This suggests that extracellular acidification reduces channel availability. Since low pH apparently also accelerated open-state inactivation, as the mean burst duration was decreased and the interburst interval was increased, these data suggest that extracellular acidification enhances both open- and closed-state inactivation, and that the two processes can coexist in Kv channels.

Voltage-clamp fluorimetry allows real-time observation of the conformational changes associated with channel gating, including those that do not result in ionic current flow (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). Changes in the emission observed from a fluorophore attached at specific residues are due to changes of its microenvironment as channels activate and inactivate (Cha and Bezanilla, 1997; Loots and Isacoff, 1998; Bezanilla, 2002). Here, we have used this technique to investigate the fate of channels at low pH by directly monitoring the effect of acidification on the channel rearrangements associated with P-type, C-type, and closed-state inactivation of Shaker channels. We show that low extracellular pH not only enhances P-type inactivation of channels, but also their progression to the C-type inactivated state, and in addition that protons stabilize an unavailable state in which channels are inactivated at rest. We demonstrate that stabilization of closed-inactivated states at low pH is largely responsible for the decrease in peak current amplitude, while stabilization of open-inactivated states is responsible for the enhanced current decay. In parallel, we have obtained single channel and gating current data that are in accord with this hypothesis. These data provide a direct demonstration that protons alter both open- and closed-state inactivation of Kv channels.

	MATERIALS AND METHODS

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Solutions
For Xenopus oocytes, Barth's medium contained (in mM) 88 NaCl, 1 KCl, 2.4 NaHCO₃, 0.82 MgSO₄, 0.33 Ca(NO₃)₂, 0.41 CaCl₂, 20 HEPES, titrated to pH 7.5 using NaOH. ND96 bath solution contained (in mM) 96 NaCl, 3 KCl, 1 MgCl₂, 0.3 CaCl₂, titrated to pH 7.5, 6.0, 5.0, or 4.0 using NaOH or HCl. HEPES (5 mM) was used as the buffer for pH 7.5 and pH 6.0 solutions, while 5 mM MES was used to buffer pH 5.0 and pH 4.0 solutions. Tetramethylrhodamine-5-maleimide (TMRM; Invitrogen) labeling of oocytes was performed in a depolarizing solution consisting of (in mM) 99 KCl, 1 MgCl₂, 2 CaCl₂, and 5 HEPES, titrated to pH 7.5 using KOH and which contained 50 µM TMRM. To record Shaker channel gating currents from tsa201 cells, the pipette contained (in mM) 140 NMG, 1 MgCl₂, 10 EGTA, 10 HEPES, titrated to pH 7.5 using HCl, and the bath solution contained (in mM) 140 NMG, 1 MgCl₂, 10 HEPES, titrated to pH 7.5 or pH 4.0 using HCl. To record Shaker single channel activity from ltk– cells, the pipette contained (in mM) 130 KCl, 10 EGTA, 1.38 MgCl₂ (free concentration, 1 mM), 4.75 CaCl₂ (free concentration, 50 nM), 10 HEPES, titrated to pH 7.4 using KOH, and the bath solution contained (in mM) 3.5 KCl, 140 NaCl, 2 CaCl₂, 1 MgCl₂, 10 HEPES or MES, titrated to pH 7.4 or pH 4.0 using NaOH or HCl. Unless stated, all chemicals were purchased from Sigma-Aldrich.

Molecular Biology and RNA Preparation
The N-terminal deletion mutant Shaker 6–46 (Shaker-IR) that is fast inactivation removed (Hoshi et al., 1991) was expressed in Xenopus oocytes using a modified pBluescript SKII expression vector (pEXO) (a gift from A. Sivaprasadarao, University of Leeds, Leeds, UK) and in ltk– and tsa201 mammalian cells using the GW1 vector (a gift from G. Panyi, University of Debrecen, Debrecen, Hungary). For oocyte fluorescence experiments, cysteine residues were introduced at specific sites (M356-R362) in the S3–S4 linker and S4 voltage sensor, and S424 in the S5-P turret for TMRM labeling and the only externally accessible cysteine residue, found in the S1–S2 linker, was replaced with a valine residue (C245V) to prevent nonspecific dye labeling. For clarity, we use the terms Shaker A359C and Shaker S424C to describe the Shaker 6–46 C245V A359C and Shaker 6–46 C245V S424C mutant channels, respectively. The T449V mutation in Shaker was used to inhibit slow inactivation (Lopez-Barneo et al., 1993) and the W434F mutation was used to permanently P-type inactivate channels (Perozo et al., 1993; Yang et al., 1997). The ILT mutant channel was a gift from E. Isacoff (University of California, Berkeley, CA) and was used to isolate the independent voltage sensor transitions from the concerted opening transition (Smith-Maxwell et al., 1998a,b). Point mutations were generated using the Stratagene Quikchange kit (Stratagene). All primers were synthesized by Sigma Genosys. All constructs were sequenced at the University of British Columbia core facility. cDNA was linearized using BstEII or HindIII (for ILT channels) and cRNA was synthesized using the mMessage mMachine T7 Ultra cRNA transcription kit (Ambion).

Oocyte Preparation and Injection
Xenopus laevis oocytes were prepared and isolated as described previously (Claydon et al., 2000). In brief, gravid frogs were terminally anesthetized, and stage V–VI oocytes were isolated and defolliculated using a combination of collagenase treatment (1 h in 1 mg/ml collagenase type 1; Sigma-Aldrich) and manual defolliculation. Oocytes were injected with 50 nl (5–10 ng) cRNA using a Drummond digital microdispenser (Fisher Scientific) and then incubated in Barth's medium at 19°C. Currents were recorded 1–7 d after injection.

Voltage-Clamp Fluorimetry
Voltage-clamp fluorimetry was performed as described previously (Claydon et al., 2006). In brief, introduced cysteine residues were labeled with 50 µM TMRM in oocyte depolarizing solution. Fluorimetry was performed using a Nikon TE300 inverted microscope with epifluorescence attachment and a 9124b Electron Tubes photomultiplier tube (PMT) module (Cairn Research). TMRM was excited by light filtered with a 525-nm band pass excitation filter and the fluorescence emission was collected using a 20x objective and filtered through a 565-nm long pass emission filter before being passed to the PMT recording module. Voltage signals from the PMT were then digitized using an Axon Digidata 1322 A/D converter and passed to a computer running pClamp9 software (Axon Instruments) to record the fluorescence emission intensity. To minimize fluorophore bleaching, a computer-controlled shutter (Uniblitz; Vincent Associates) was opened shortly before voltage-clamp pulses were applied. Fluorescence signals were filtered at 1 kHz with a sampling frequency of 20 kHz (when the pulse duration was 100 ms), 10 kHz (when the pulse duration was 7 s), or 2 kHz (when the pulse duration was 42 s). Traces were not averaged, except for those recorded during 100-ms pulses, which represent the average of five sweeps. To account for the bleaching of the fluorescence signal during 7- or 42-s pulses, fluorescence recorded for 7 or 42 s at the holding potential of –80 mV was subtracted. Simultaneous voltage clamp of the oocyte and acquisition of the current and voltage signals was achieved using the two-microelectrode voltage-clamp technique with a Warner Instruments OC-725C amplifier, Axon Digidata 1322, and pClamp9 software. Microelectrodes were filled with 3 M KCl and had a resistance of 0.2–0.5 M.

Unlabeled wild-type Shaker channels inactivate with a time constant of 5.7 ± 0.7 s, and unlabeled Shaker A359C and S424C mutant channels inactivate with time constants of 5.6 ± 0.5 and 1.5 ± 0.2 s, respectively. This suggests that the introduction of a cysteine residue at S424, but not A359, speeds inactivation significantly. Despite this change, we believe that both the inactivation process and the nature of the fluorescence changes are qualitatively similar in their fast and slow components. First, we have shown that low pH accelerates inactivation of wild-type, A359C, and S424C channels similarly (9.4-, 6.9-, and 5.7-fold, respectively; the time constants of inactivation were reduced to 0.61 ± 0.11 s, 0.81 ± 0.18 s, and 0.25 ± 0.05 s, respectively). As well, other interventions that are known to modify inactivation such as raised external K⁺, the mutation C462A in S6, and also low pH (Loots and Isacoff, 1998) result in consistent changes in ionic current and fluorescence measurements. This suggests that the underlying processes being examined are not seriously perturbed in these two cysteine-substituted channels. TMRM-labeled Shaker A359C and S424C channels inactivate with time constants of 2.7 ± 0.2 and 2.0 ± 0.3 s, respectively. Loots and Isacoff (1998) reported an inactivation time constant for TMRM-labeled S424C of 2.5 s, but a value for inactivation of ionic current in A359C mutant channels has not been reported previously. Our data suggest that TMRM labeling of A359C, but not S424C, speeds inactivation twofold (Table I). However, as suggested above, inactivation in TMRM-labeled A359C channels remains modifiable by low pH (the time constant of inactivation is reduced 4.6-fold to 0.58 ± 0.08 s at pH 4.0) and the overall effect is relatively unchanged. This also holds true for the pH sensitivity of TMRM-labeled S424C channels, which was somewhat greater than in unlabeled channels.

Data Analysis
G-V curves were derived using the normalized chord conductance, which was calculated by dividing the maximum current during a depolarizing step by the driving force derived from the K⁺ equilibrium potential (the internal K⁺ concentration was assumed to be 99 mM). G-V, F-V, and Q-V curves were fitted with a single Boltzmann function:

(1)

where y is the conductance normalized with respect to the maximal conductance, V_1/2 is the half-activation potential, V is the test voltage, and k is the slope factor. Data throughout the text and figures are shown as mean ± SEM.

	RESULTS

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Acidic pH Reduces the Conductance of Shaker 6–46 Channels
Shaker 6–46 currents are inhibited by extracellular protons (pK_a 4.7–5.0) and this has been attributed to enhanced P/C-type open-channel inactivation at low pH (Perez-Cornejo, 1999; Starkus et al., 2003) that accumulates during repetitive stimulation (Starkus et al., 2003). However, we found that acidic extracellular pH reduced current amplitude even when the acceleration of inactivation during the depolarization was minimal, suggesting that protons also reduce the macroscopic conductance. This is shown in Fig. 1 A in the comparison of conductance–voltage relations at different external pH, which were constructed from currents recorded in response to short voltage-clamp pulses (100 ms) and with long interpulse intervals (10 s) to avoid cumulative inactivation. In addition to a rightward shift of the voltage dependence of channel opening that is attributed to a surface charge-screening effect (Deutsch et al., 1989; Hille, 2001; Kehl et al., 2002; Trapani and Korn, 2003), the maximal conductance was reduced at pH 5.0 by 21 ± 7% and at pH 4.0 by 47 ± 5% (n = 3).

View larger version (20K): [in this window] [in a new window] [Download PPT slide]   Figure 1. Low pH reduces the conductance of Shaker channels. (A) G-V relations of Shaker 6–46 channels recorded with the indicated external pH (n = 3). Conductance was calculated from currents recorded during 100-ms voltage pulses applied from –80 to +100 mV at 10-s intervals in 10-mV increments (holding potential –80 mV). (B–E) Typical ionic currents (B and C) and fluorescence signals (D and E) recorded from Shaker A359C channels in response to 100-ms voltage pulses applied from –80 to +100 mV at 10-s intervals (holding potential –80 mV) at pH 7.5 and pH 4.0. Although pulses were applied in 10-mV increments, only selected pulses are shown for clarity. Current and fluorescence traces at pH 4.0 (C and E) are shown at more depolarized potentials than in B and D to account for the charge screening effect of protons. Contributions of the fast and slow phases to the overall fluorescence signal were measured from biexponential fits of the fluorescence traces at +60 mV for pH 7.5 and at +100 mV for pH 4.0. Arrows and percentages in D and E show the contribution of the slow phase at pH 7.5 and pH 4.0. The mean slow phase contribution was 14 ± 2% at pH 7.5 and 71 ± 3% at pH 4.0 (n = 8).

Data in Fig. 1 (B–E) show ionic current traces and fluorescence signals recorded from Shaker A359C channels with an extracellular pH of either 7.5 or 4.0. TMRM labeling of A359C has previously been shown to faithfully report local environmental changes associated with voltage sensor movement in response to membrane depolarization (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). At pH 7.5, there was no obvious channel opening at the holding potential of –80 mV in Fig. 1 B and no voltage sensor movement in Fig. 1 D. On depolarization to –60 mV, despite the low open probability, there was a significant fluorescence report consistent with voltage sensor movement (Fig. 1 D). Further depolarization to +60 mV induced robust ionic currents that activated rapidly and showed only minimal decay (Fig. 1 B). This was reflected in the fluorescence signals (Fig. 1 D), which show a large fluorescence deflection from Shaker A359C channels with a fast phase of fluorescence quenching that accounted for 86 ± 2% (n = 8) of the total fluorescence deflection and occurred with a time course similar to that for channel activation. Biexponential fits to the fluorescence relaxation showed that this rapid component was followed by a slow phase that accounted for 14 ± 2% of the deflection and was associated with channel inactivation (Claydon et al., 2006).

At pH 4.0, peak ionic current was reduced by 57 ± 4% (Fig. 1 C; n = 21) even when charge screening was accounted for by the application of a depolarizing pulse to +100 mV, at which open probability was maximal (Fig. 1 A). However, the corresponding fluorescence signals in Fig. 1 E show that the fluorophore continues to report the same extent of environmental change (although with altered kinetics; see below), which suggests that although a proportion of the channels does not conduct at acidic pH, voltage sensor movement is preserved. The mean peak amplitude of the fluorescence deflection at pH 4.0 was 116 ± 11% of that at pH 7.5 (n = 7; t test, not significantly different).

Another measure of voltage sensor movement can be obtained from gating current measurements. We recorded gating currents from wild-type Shaker channels at pH 7.5 and pH 4.0 (Fig. 2) and a typical family of records obtained at pH 7.5 during 20-ms voltage-clamp pulses ranging from –80 to +60 mV (the holding potential was –80 mV) is shown in Fig. 2 A (current during the first 6 ms of the pulse is shown to highlight on-gating events). In Fig. 2 B, the integrals of the on-gating currents recorded at pH 7.5 and pH 4.0 in the same cell are plotted. These show that acidic pH shifted the voltage dependence of charge movement to more depolarized potentials, but did not significantly alter the total gating charge movement. This is shown more clearly by the mean Q-V relations in Fig. 2 C. Acidic pH shifted the V_1/2 of the Q-V relation by +35 mV from –27.9 ± 0.4 mV (k was 9.5 ± 0.4 mV) at pH 7.5 to +5.9 ± 1.2 mV (k was 12.4 ± 1.0 mV) at pH 4.0, without altering the total gating charge, which suggests that the number of activatable channels (i.e., the number of channels in which the voltage sensors are available to move) is not changed.

View larger version (12K): [in this window] [in a new window] [Download PPT slide]   Figure 2. Acidic pH does not alter total gating charge movement in Shaker channels. (A) Typical wild-type Shaker 6–46 (Shaker A359) gating current records obtained from tsa201 cells in the whole-cell patch clamp configuration during 20-ms voltage-clamp pulses from –80 to +60 mV (in 10-mV increments) from a holding potential of –80 mV. Currents during every other pulse are shown for clarity and only the first 6 ms of the depolarizing pulses are shown to highlight the on-gating current. (B) Typical integrals of on-gating currents, such as those in A, recorded in the same cell under control conditions (pH 7.5) and at pH 4.0. Acidic pH did not alter the total gating charge movement in these channels. (C) Mean Q-V relationships recorded at pH 7.5 and pH 4.0 (n = 2–4). At pH 4.0, the Q-V relation was shifted by +35 mV from –27.9 ± 0.4 mV at pH 7.5 to +5.9 ± 1.2 mV.

Acidic pH Accelerates the Inactivation Report from Shaker A359C Channels
The effect of acidic pH on Shaker A359C channel inactivation during 7-s depolarizations to +60 mV is shown in Fig. 3 (see also Table I). The fluorescence signals in Fig. 3 B show a clear slow phase that corresponds to the inactivation of ionic current in panel A. At pH 7.5, ionic current decayed with a of 2.7 ± 0.2 s (Fig. 3 A; n = 14) and the slow phase of fluorescence decayed with a similar of 2.4 ± 0.3 s, and contributed 53 ± 4% to the total signal amplitude (Fig. 3 B; n = 14). Reducing the extracellular pH to 5.0 had only a minor effect on ionic current and the fluorescence signal from Shaker A359C channels (Figs. 3, A and B). However, pH 4.0 both reduced the peak current amplitude and enhanced the decay of the remaining current (Fig. 3 A; was 579 ± 84 ms, n = 11). The fluorescence signals in Fig. 3 B, which were recorded simultaneously, show that at pH 4.0 both the contribution and the decay of the slow deflection were enhanced; the contribution of the slow phase was increased to 76 ± 3% and the of the slow decay was reduced to 1.2 ± 0.1 s (n = 14). These data confirm the previous reports that low pH enhances inactivation of Shaker channels (Perez-Cornejo, 1999; Starkus et al., 2003) and suggest that acidic pH enhances the conformational rearrangements associated with inactivation, which are reported by a fluorophore at the outer end of the voltage sensor.

View larger version (10K): [in this window] [in a new window] [Download PPT slide]   Figure 3. Acidic pH enhances the inactivation report from Shaker A359C. Typical ionic currents (A) and fluorescence signals (B) recorded from Shaker A359C channels during 7-s voltage-clamp pulses to +60 mV at the indicated external pH (holding potential of –80 mV). (C) Typical ionic currents and fluorescence signals recorded at pH 7.5 and pH 4.0 from TMRM-labeled channels in which a cysteine residue was not engineered at position A359 (Shaker A359). These channels lack an externally labelable cysteine residue (the only external cysteine, in the S1–S2 linker [C245], has been removed) and act as a control demonstrating the lack of a voltage-dependent fluorescence deflection in the absence of an introduced cysteine and also the pH independence of emission of the TMRM dye.

To demonstrate the specificity of the response of the fluorophore attached at position A359 to changes in external pH, we compared the effect of changing the pH on TMRM-labeled Shaker channels lacking an externally labelable cysteine residue (Shaker A359; Fig. 3 C and Table I). The decay of ionic current was accelerated from 3.9 ± 0.5 s at pH 7.5 to 0.49 ± 0.07 s at pH 4.0 (n = 3–4; Table I; these values are not significantly different from wild type in the absence of TMRM). The fluorescence signals from Shaker A359 demonstrate that there are no voltage-dependent conformational changes reported by TMRM that labels nonspecific sites in endogenous membrane proteins. Furthermore, the fluorescence signal is not altered by changes of the extracellular pH. These results emphasize the ability of site-specific attachment of TMRM to detect gating-induced rearrangements of channel conformation, and in addition, confirm that the emission from TMRM is pH insensitive (Cha and Bezanilla, 1998; see also Molecular Probes website at http://probes.invitrogen.com).

Detection of the pH-induced Acceleration of Inactivation by the Voltage Sensor Is Site Specific
To understand which structural domains are involved in the pH-induced changes in the fluorescence signal from TMRM attached at A359C and the local rearrangements that they represent, we investigated the effect of acidic pH on the fluorescence signal from TMRM attached at each position from M356C in the S3–S4 linker to the outermost arginine, R362C, in the S4 domain. Fluorescence signals recorded during 7-s depolarizing pulses to +60 mV from each of these mutant channels at either pH 7.5 or pH 4.0 extracellular solution are shown in Fig. 4. Each site reported robust fluorescence signals on depolarization with the exception of I360C and L361C, from which only baseline signals could be detected (unpublished data). This lack of signal was unexpected given the fluorescence report from these residues described previously (Loots and Isacoff, 2000). However, we were unable to detect voltage-dependent signals in any of 17 oocytes, which displayed large ionic currents. Close examination of the effect of acidic pH on the fluorescence signal from each mutant channel reveals that only A359C and R362C reported pH-dependent changes. With TMRM attached at R362C, the of the slow phase was reduced from 2.6 ± 0.4 s at pH 7.5 to 1.4 ± 0.2 s at pH 4.0 (n = 3; P < 0.05, paired t test). Since A359C and R362C are likely positioned on the same side of the S4 helix, this suggests that this face senses the rearrangements associated with the pH modulation of inactivation.

View larger version (7K): [in this window] [in a new window] [Download PPT slide]   Figure 4. Voltage sensor detection of pH-induced enhancement of inactivation is site specific. Typical fluorescence signals from TMRM attached at different sites in the S4 voltage sensor and S3–S4 linker from M356C to R362C recorded during a 7-s pulse to +60 mV at pH 7.5 and pH 4.0 (holding potential –80 mV). Robust fluorescence deflections were recorded from each site in the scan with the exception of I360C and L361C, from which we were unable to record voltage-dependent fluorescence signals. Only A359C and R362C report the pH-induced enhancement of inactivation.

View larger version (10K): [in this window] [in a new window] [Download PPT slide]   Figure 5. Two effects of acidic pH are detected from TMRM attached within the pore. Typical ionic currents (A) and fluorescence signals (B) recorded from Shaker S424C channels during 42-s voltage-clamp pulses to +60 mV at the indicated external pH (holding potential –80 mV). Acidic pH decreased peak and accelerated inactivation of the ionic current, and this was detected in the fluorescence signals as a decrease in the peak fluorescence amplitude and an enhancement of the slow phase of the remaining fluorescence deflection. (C) Shaker S424C channel fluorescence deflections from B scaled to the maximum signal amplitude to highlight the pH-induced enhancement of the slow phase of fluorescence deflection.

To understand why the macroscopic conductance and the fluorescence report from TMRM at S424C was reduced with acidic pH (Fig. 5), while the report from A359C was not (Figs. 1 and 3), we measured Shaker-IR single channel currents at pH 7.4 and pH 4.0 (Fig. 6), since the reduced report from S424C may reflect an effect of protons on channel open probability or the single channel current amplitude. Channel activity was assessed during 500-ms voltage-clamp pulses to +100 mV from outside-out patches (the pulse interval was 15 s). At pH 7.4, the single channel current amplitude (at +100 mV) was 2.7 ± 0.3 pA and the representative traces show high channel availability. We calculated the availability by determining the proportion of sweeps showing channel activity during the first 50 ms of the pulse, and the average availability from 11 patches was 0.87 at pH 7.4. Application of pH 4.0 to the same patch (Fig. 6) markedly reduced availability, resulting in a number of null sweeps (the average availability from five patches was 0.14), but did not alter the single channel current amplitude (2.7 ± 0.3 pA). These data suggest that the loss of fluorescence observed in Fig. 5 is not due to reduced single channel conductance, but rather reduced channel availability.

View larger version (26K): [in this window] [in a new window] [Download PPT slide]   Figure 6. Acidic pH decreases open probability without altering single channel current amplitude. Typical recordings of single channel events obtained from outside-out patches of ltk– cells expressing Shaker 6–46 channels during consecutive 500-ms voltage-clamp pulses to +100 mV (pulse interval 15 s) from a holding potential of –80 mV (the voltage protocol is shown). In this example the patch contains a single channel. Channel activity from the same patch was recorded during control conditions (pH 7.4) and at pH 4.0. Dashed lines mark the zero current level. Acidic pH markedly reduced channel availability (calculated by determining the proportion of sweeps showing channel activity during the first 50 ms of the pulse) from 0.87 at pH 7.4 to 0.14 at pH 4.0 without effect on the single channel current amplitude. At +100 mV, the single channel current amplitude was 2.7 ± 0.3 pA at pH 7.4 and 2.7 ± 0.3 pA at pH 4.0 (data were collected from 11 patches at pH 7.4 and 5 patches at pH 4.0). The 84% reduction in channel availability (from 0.87 to 0.14) was greater than the reduction of macroscopic conductance measured from Shaker channels expressed in Xenopus oocytes (Fig. 1), but similar to the decrease in peak macroscopic current measured from Shaker channels expressed in mammalian cells at pH 4.0 (91 ± 8%; n = 5). This difference between expression systems is most likely due to incomplete exchange of solution over the entirety of the large invaginated oocyte membrane.

View larger version (11K): [in this window] [in a new window] [Download PPT slide]   Figure 7. The loss of fluorescence at low pH is similar to the reduction of peak current amplitude. (A) G-V relations of Shaker S424C channels recorded with the indicated external pH (n = 3). Using Eq. 1 (the intracellular K+ concentration was assumed to be 99 mM), conductance was calculated from currents recorded during 100-ms voltage pulses applied from –80 to +100 mV at 10-s intervals in 10-mV increments (holding potential –80 mV). (B) F-V relations of the Shaker S424C channels recorded at the indicated pH during 42-s pulses from the same group of oocytes as A (n = 3). All fluorescence amplitudes were normalized to the fluorescence amplitude at +100 mV with pH 7.5. Lines represent no mathematical significance and are simply to guide the eye. The loss of fluorescence at low pH was similar to the decrease in conductance at all potentials. (C) Plot of the dependence on the external pH of Shaker S424C channel fluorescence amplitude at +60 mV or conductance at +60 mV (n = 2–7). Fluorescence and conductance values were normalized to those at pH 7.5. Conductance values were calculated from peak currents recorded at the same time as fluorescence deflections during 42-s pulses so as to directly compare the decrease in conductance with the loss of fluorescence from simultaneous measurements from the same oocyte. Data were fitted with a standard Hill equation (assuming a Hill coefficient, n, of 1). pKa values were 5.1 and 5.2 for the loss of fluorescence and reduction of conductance, respectively.

The data in Figs. 5–7 demonstrate that acidic pH has two effects: an acceleration of the slow phase of fluorescence from Shaker S424C that represents the enhancement of inactivation upon depolarization, and a loss of the fluorescence report from S424C that, in the light of the observed reduction in single channel open probability, suggests a reduction of channel availability and consequently the peak current amplitude. The pH- induced loss of fluorescence signal is of particular interest because it contrasts with the lack of effect of acidic pH on the fluorescence amplitude recorded from Shaker A359C channels (Fig. 1 F and Fig. 3 B) and allows a quantitative correlation to be made between the fluorescence loss and conductance decrease at low pH.

Acidic pH Enhances Rearrangements Associated with P- and C-type Inactivation
P/C-type inactivation involves two sequential processes, with a concerted closure of the pore resulting in the collapse of conductance (P-type inactivation) followed by a stabilization of the voltage sensor in its activated conformation (C-type inactivation), which reports as an immobilization of the return of gating charge on repolarization (De Biasi et al., 1993; Olcese et al., 1997; Yang et al., 1997; Loots and Isacoff, 1998). Channels with the mutation W434F are thought to be permanently P-type inactivated (Perozo et al., 1993; Yang et al., 1997), and the fluorescence signal from TMRM attached at S424C in these channels reports well on activation, as if P-type inactivation moves the fluorophore into closer proximity with the voltage sensor and allows it to track the transition of channels to a W434F-like inactivated state (Loots and Isacoff, 1998). This idea is consistent with the close proximity of the S5-P linker with S4 (of an adjacent subunit) in the Kv1.2 channel crystal structure (Long et al., 2005). Since, in the present study, acidic pH accelerates the slow fluorescence report from S424C, this suggests that protons enhance P-type inactivation and we therefore used a protocol designed to monitor the effect of pH on the ability of TMRM attached at S424C to detect voltage sensor movement. Fig. 8 A shows ionic current and fluorescence signals recorded from Shaker S424C TMRM-labeled channels during a 2-s test pulse to +60 mV applied either 200 ms, 12 s, or 42 s after a 5-s conditioning pulse to +60 mV. When the interval between pulses was short (200 ms), only a small percentage of channels recovered from inactivation and so the fluorescence signal during the test pulse reports mostly from inactivated channels. In this case, the amplitude of the fast phase of the fluorescence signal was increased from 26 ± 3% in the conditioning pulse to 60 ± 3% during the test pulse (n = 6; P < 0.01, paired t test), suggesting that many channels became P-type inactivated during the conditioning pulse as the fluorophore now reports activation more clearly. With a 42-s interval that allowed for more complete recovery from inactivation, the fluorescence signal during the test pulse recovered its slow phase and the fast phase diminished back to 17 ± 7%. Fig. 8 B shows the effect of acidic pH on the progression of channels to the P-type inactivated state. The double pulse protocol shows that the fast phase dominated during the test pulse at pH 5.0, suggesting that the majority of the channels sense activation well and are therefore P-type inactivated. The contribution of the fast phase in the test pulse after a 200-ms interval at pH 5.0 was 82 ± 4%, which is significantly greater (n = 5; P < 0.01, paired t test) than the 60 ± 3% seen at pH 7.5 (Fig. 8 A). These data demonstrate that the enhancement of current decay at acidic pH is a result of enhanced P-type inactivation.

View larger version (11K): [in this window] [in a new window] [Download PPT slide]   Figure 8. Low pH enhances P-type inactivation. Typical ionic currents and fluorescence signals recorded at pH 7.5 (A) and pH 5.0 (B) from Shaker S424C channels during a 2-s test pulse to +60 mV applied either 200 ms, 12 s, or 42 s after a 5-s conditioning pulse to +60 mV (holding potential –80 mV). Similar records were obtained from five other oocytes.

View larger version (8K): [in this window] [in a new window] [Download PPT slide]   Figure 9. Low pH enhances C-type inactivation. (A) Typical fluorescence signals recorded from the same cell expressing Shaker S424C W434F channels during 7-s depolarizing pulses to +60 mV (applied after at least 3 min at the holding potential of –80 mV) at pH 7.5, pH 5.0, and on return to pH 7.5. Oocytes were held at –80 mV for 3 min between measurements. The W434F mutation permanently P-type inactivates channels enabling observation of C-type inactivation rearrangements during depolarizing pulses. The amplitude of the decaying component of fluorescence was approximately twofold larger at pH 5.0 than at pH 7.5. The dotted lines highlight the extent of the decay. The decay of fluorescence was biexponential with values of 21 ± 1 ms and 1.8 ± 0.1 s at pH 7.5, and 21 ± 1 ms and 0.36 ± 0.03 s at pH 5.0 (n = 3; P < 0.001, paired t test, when compared with pH 7.5). (B) Plot of the voltage dependence of the amplitude of the decaying component of fluorescence that reflects C-type inactivation (n = 3). The amplitude of the decaying component was measured at each potential and normalized to that at +100 mV (F decay). Also plotted is the G-V relationship of Shaker A359C channels and F-V relationship of Shaker A359C W434F channels for comparison of the voltage dependence of pore opening and voltage sensor movement, respectively (n = 5–21).

View larger version (7K): [in this window] [in a new window] [Download PPT slide]   Figure 10. Inhibition of inactivation by raising external K+ rescues the loss of fluorescence at low pH. (A–F) Typical fluorescence signals recorded from the same oocyte expressing Shaker S424C channels during 42-s depolarizing pulses to +60 mV during the indicated manipulations of the external K+ and proton concentration. After control fluorescence recordings were obtained with 3 mM K+ and pH 7.5 (A), the pH was reduced to pH 5.0 (with 3 mM K+) to demonstrate the decrease of fluorescence (B). Fluorescence signals were then recorded on return to pH 7.5 with 3 mM K+ (C) before application of solution with 99 mM K+ at pH 7.5 (D). The fluorescence was then measured with 99 mM K+ and pH 5.0 (E), before the oocyte was washed with control solution (3 mM K+ and pH 7.5) again (F). Similar records were obtained from three other oocytes.

View larger version (9K): [in this window] [in a new window] [Download PPT slide]   Figure 11. Inhibition of inactivation by the mutation T449V rescues the loss of fluorescence at low pH. Typical fluorescence signals recorded from Shaker S424C T449V channels during 7-s depolarizing pulses to +60 mV at pH 7.5 and pH 5.0 (holding potential –80 mV). Inhibition of inactivation prevented the loss of fluorescence at low pH.

View larger version (17K): [in this window] [in a new window] [Download PPT slide]   Figure 12. Acidic pH induces conformational changes in closed channels that are associated with inactivation. Typical fluorescence signals recorded from Shaker ILT S424C channels during 100-ms voltage pulses from –80 to 0 mV at pH 7.5 (A) and from –80 to +80 mV at pH 5.0 (B). Although pulses were applied in 10-mV increments, only every other pulse is shown for clarity. A voltage pulse to 0 mV at pH 7.5 or to +80 mV at pH 5.0 (shift in voltage dependence is due to surface charge screening by protons) evokes maximum voltage sensor movement without any channel opening. Reduction of the extracellular pH induced conformational changes in closed channels that are consistent with inactivation. (C) Mean F-V relationships of the fluorescence recorded from closed channels at pH 7.5 and pH 5.0 from Shaker ILT S424C channels (n = 4), and from Shaker ILT A359C W434F channels (n = 4). The V1/2 and k values were –37.1 ± 0.8 mV and 16.1 ± 0.7 mV for Shaker ILT S424C channels at pH 5.0, and –83.0 ± 0.4 mV and 15.7 ± 0.4 mV for Shaker ILT A359C W434F channels at pH 7.5, respectively. (D) Mean -V relationships for the fast phase (fitted with a single exponential) of the fluorescence deflection from Shaker ILT S424C channels at pH 5.0, and from Shaker ILT A359C W434F channels (n = 4). The voltage dependence and kinetics of the fluorescence report from Shaker ILT S424C channels was similar to that of voltage sensor movement reported by Shaker A359C W434F.

View larger version (11K): [in this window] [in a new window] [Download PPT slide]   Figure 13. Demonstration of pH-induced closed-state rearrangements. Typical diary plots of the effect of changing the pH on the fluorescence emission from Shaker ILT S424C (A) or Shaker A359 (C, as a control) channels held continuously at –80 mV. To minimize bleaching of the fluorophore, fluorescence was sampled every second (although only every fifth recording is shown for clarity) by opening the shutter for 100 ms. Low pH induced rearrangements at –80 mV that were detected by TMRM at S424C. (B) Plot of the dependence of the closed-state fluorescence changes on the external pH (n = 3). The relative fluorescence change from that at pH 7.5 is plotted. Data were fitted with a standard Hill equation (assuming a Hill coefficient, n, of 1). The pKa for the closed-state fluorescence change was pH 5.9. The value at pH 7.5 represents the mean of all pre- and post-treatment conditions since the order of pH changes was not necessarily consistent.

	DISCUSSION

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It is interesting that the large fluorescence deflection observed with Shaker ILT S424C (Fig. 12) was not observed in Shaker S424C channels at low pH (Fig. 5). Those channels that are closed inactivated should report a rapid fluorescence deflection on depolarization, but this is not evident. This may be because this report is masked by the robust signal from open and inactivating Shaker S424C channels at low pH. Indeed, Fig. 9 A shows that permanently inactivated Shaker S424C channels produce only a small fluorescence deflection (see also Fig. 4 C in Loots and Isacoff, 1998). It appears that the ILT mutation can enhance the report of activation in channels that are closed-state inactivated in some way that is not fully understood at the present time.

A positive correlation between the rate of depolarization-induced P/C-type inactivation and the prevalence of closed-state inactivation has been reported previously (Lopez-Barneo et al., 1993). These authors showed that reducing external K⁺ reduced channel availability (i.e., decreased the proportion of channels able to conduct ions on depolarization) more significantly in mutant Shaker channels that displayed accelerated depolarization-induced inactivation. This conclusion is also supported by the demonstration that the loss of fluorescence and the decrease of macroscopic conductance share a very similar pH dependence with pKa values of 5.1 and 5.2, respectively (Fig. 7), and is consistent with the observation that low pH markedly reduced the open probability of Shaker single channels (Fig. 6). Similar findings are reported for the effect of acidic pH on Kv1.5 channels in which single channels switch between available and unavailable states (Kwan et al., 2006). This predicts the loss of the macroscopic fluorescence signal and maximal conductance that we observe since these measurements represent the proportion of channels that are in the unavailable state. Our fluorescence data from Shaker channels demonstrate that the pKa for the transition to the unavailable closed-inactivated state is pH 5.1–5.9 (Fig. 7 C and Fig. 13 B). The pKa values for the stabilization of closed-state inactivation and loss of conductance (pH 5.2, Fig. 7 C) are similar to previously reported pKa values, pH 4.7–5.0, for the effect of protons on P/C-type inactivation from the open state (Perez-Cornejo, 1999; Starkus et al., 2003). In Kv1.5 channels, the pKa values for the inhibition of maximal conductance and the acceleration of inactivation by external acidification only differ by 0.3 pH units (pKa = pH 6.2 and pH 6.5, respectively; unpublished data). This suggests that the effects of acidic pH on closed- and open-state inactivation may involve the same mechanism.

An argument against this idea is the differential sensitivity of pH-induced closed- and open-state inactivation to external K⁺. For example, raising external K⁺ from 3 to 99 mM completely reversed the loss of the fluorescence signal caused by acidic pH, suggesting that 99 mM K⁺ prevents closed-state inactivation entirely (Fig. 10). However, the report of P/C-type inactivation was only slowed approximately twofold (from 2.0 ± 0.6 to 3.6 ± 0.4 s; n = 3) and not completely prevented (Fig. 10). Differential sensitivities of P/C-type inactivation and the inhibition of maximal conductance by acidic pH to external K⁺ are also evident in Kv1.5 channels (Fedida et al., 1999; Zhang et al., 2005). However, this difference is probably due to the fact that the K⁺-sensitive site in the pore that mediates inactivation is flooded by efflux of internal K⁺ in open channels such that it is relatively insensitive to changes in external K⁺, whereas the absence of K⁺ efflux in closed channels reveals the sensitivity of inactivation to the external K⁺ concentration, as is the case with depolarization-induced inactivation in the presence of an internal pore blocker or with the removal of internal K⁺ (Baukrowitz and Yellen, 1995).

Enhancement of P-type Inactivation at Low pH
It is well established that acidic extracellular pH enhances the rate of inactivation upon depolarization in a number of Kv channels, such as Shaker, Kv1.4, and Kv1.5, although their sensitivity to protons differs (Steidl and Yool, 1999; Claydon et al., 2000; Kehl et al., 2002; Starkus et al., 2003). Our fluorescence studies enable direct observation of the rearrangements associated with the collapse of the pore during P-type inactivation from the report of TMRM attached either within the pore at S424C or at the top of the voltage sensor at A359C. Acidic pH enhanced the slow phase of the fluorescence report from TMRM attached at A359C and at S424C that reports the dynamic structural changes associated with P-type inactivation (Fig. 3 B and Fig. 5 B). In addition, the experiments in Fig. 6 suggest that a greater proportion of channels make the transition at rest to the W434F-like P-type inactivated state at pH 5.0 than at pH 7.5. These findings are consistent with the enhanced decay of current seen in Fig. 3 A and Fig. 5 A and also with data from previous studies during exposure to acidic pH (Claydon et al., 2002; Kehl et al., 2002; Starkus et al., 2003; Zhang et al., 2003), as well as the enhanced decay of fluorescence that was previously observed from TMRM attached at S424C (Loots and Isacoff, 1998).

Enhancement of C-type Inactivation at Low pH
W434F mutant channels are thought to be permanently P-type inactivated (Perozo et al., 1993; Yang et al., 1997), and therefore transitions to the C-type inactivated state should be observable on depolarization from a fluorophore placed in the outer pore. Our fluorescence records from TMRM attached at S424C in W434F mutant channels (Fig. 9 A) showed a large fast deflection consistent with previous fluorescence observations in this channel (Loots and Isacoff, 1998) and also the notion that the fluorophore in P-type inactivated channels occupies a position that senses the movement of the voltage sensors very well (Loots and Isacoff, 1998). However, in addition to the rapid fluorescence deflection, we observed a secondary fluorescence deflection that most likely represents the progression of channels to the C-type inactivated state, and this was enhanced during acidic pH (Fig. 9, A and B). The C-type inactivation fluorescence transition was best fit with a double exponential function, and although the mechanistic basis for this is uncertain, only the slower time constant was reduced by acidic pH. The voltage dependence of the total amplitude of the fluorescence deflection followed that expected for channel opening (Fig. 9 C). This suggests that, although these channels are permanently P-type inactivated, activation of the voltage sensors is not sufficient to cause C-type inactivation, and that transition to the C-type inactivated state can only occur once the intracellular gate opens. Furthermore, this suggests that, unlike P-type inactivation, C-type inactivation cannot occur from resting closed states.

Although we have demonstrated a clear secondary fluorescence deflection that probably reflects C-type inactivation, it was not evident in the previous report of fluorescence from S424C in W434F mutant channels (Loots and Isacoff, 1998). This could be due to an additional mutation (C462A) made in those studies, which enhanced the rate of inactivation 20-fold (Loots and Isacoff, 1998) and perhaps made the transition to C-type inactivation too rapid to resolve.

A Discrete Site in S4 Reports the pH-induced Alteration of Inactivation
It has previously been suggested that the S4 domain moves close to, and even interacts with, the S5-P linker in an adjacent subunit of the channel (Blaustein et al., 2000; Elinder et al., 2001; Gandhi et al., 2000; Larsson and Elinder, 2000; Li-Smerin et al., 2000; Loots and Isacoff, 2000; Ortega-Saenz et al., 2000; Lainè et al., 2003; Long et al., 2005) and that TMRM attached at A359C at the top of the voltage sensor reports inactivation gating rearrangements (Loots and Isacoff, 1998, 2000; Gandhi et al., 2000; Long et al., 2005; Claydon et al., 2006). It is clear from the present study that A359C and R362C are unique amongst sites in the S3–S4 linker and at the top of S4 in their ability to sense the pH- induced enhancement of inactivation (Fig. 4). In a previous comprehensive scan of the extracellular domains of the Shaker channel (Gandhi et al., 2000) it was demonstrated that all sites from M356C to R362C report on inactivation of the pore and that TMRM at sites within the outer portion of the helical S4 domain can sense inactivation rearrangements regardless of their orientation (Loots and Isacoff, 2000). Our results are generally consistent with these (except that we could not record deflections from either I360C or L361C); however, the data in Fig. 4 suggest that only two of these sites, which are adjacent to each other on the same side of the S4 -helix that faces the pore, sense the pH-induced changes (although we cannot exclude that I360C and L361C might sense such changes). It is unclear why we were unable to record fluorescence deflections from I360C and L361C (even with oocytes expressing >200 µA of current) that were reported by Loots and Isacoff (2000). Since these authors suggest that these two sites report only on inactivation (because the fluorescence signal is dominated by a slow component and there is little evidence of a fast component on the same time scale as activation), it is possible that their inclusion of the mutation C462A is the reason for the discrepancy. This mutation is known to speed inactivation 20-fold (Loots and Isacoff, 1998) and the report of inactivation in this background would therefore be expected to be much more pronounced.

Our data suggest either that the inactivation gating rearrangements that are induced at low pH are localized so that they are only sensed by this region or that protons alter the interaction of S4 with the pore and, as a consequence, the influence of inactivation on TMRM attached in S4. Further studies are clearly required to address the effect of pH on the dynamic interaction of S4 with the pore.

It has been suggested that the extent of fluorescence quenching of TMRM attached at A359C is increased with acidic pH (Cha and Bezanilla, 1997; Sorensen et al., 2000), an effect that was attributed to the protonation of a titratable group within the S3–S4 linker. In the present study, we observed a trend for the fluorescence deflection to increase in amplitude during application of acidic pH, although the data did not reach statistical significance (Fig. 1, D and E). However, the key observation was that the fluorescence amplitude from Shaker A359C was not reduced during acidic pH (Fig. 1, D and E). The lack of reduction of the signal strongly suggests that the total gating charge movement was not altered by external protons and is consistent with the observation that acidic pH did not alter the amplitude of on-gating charge movement (Fig. 2 C). Clearly, the kinetics of the report of channel gating are dramatically altered during acidic pH (Fig. 1, D and E), but we do not interpret this as a reduction in the gating charge movement, given that the total signal amplitude is unchanged, but rather that the enhancement of the inactivation report masks the report of activation. This suggests that although the pore reports a loss of channel availability at low pH, the number of activatable channels is not reduced.

Other Channels and Other Mechanisms
In Kv1.5 channels, although the pKa for the inhibition by protons is pH 6.2 with 2–5 mM external K⁺ (Steidl and Yool, 1999; Kehl et al., 2002), lowering the pH to 5.5 results in a loss of conductance of channels in which the histidine, which acts as a pH sensor in the outer pore, has been mutated to glutamine (H463Q; Kehl et al., 2002). This suggests that there are two titratable sites in the Kv1.5 pore, one that has a pKa of pH 6.2 and, when protonated, enhances inactivation, and a second that has a lower pKa, which, when protonated, causes a loss of macroscopic conductance. The Shaker channel lacks the histidine pH sensor, and current is inhibited with a pKa of pH 4.0–5.0 (Perez-Cornejo, 1999; Starkus et al., 2003; see also Fig. 1 A), leading to the suggestion that D447 in the outer pore may act as the pH sensor (Starkus et al., 2003). Since this residue is conserved in Kv1.5 channels, this site may also act as the low range pH sensor that regulates availability in these channels. Consistent with this idea, mutations of Shaker D447 accelerate inactivation (D447E) or alter current level (D447N) (Molina et al., 1998).

It is known that Kv channels, once inactivated, become significantly more permeable to Na⁺ (Starkus et al., 1997; Wang et al., 2000), and this has been used as another way to examine transitions between inactivated states at low pH (Zhang et al., 2003). Many of the observations made on Na⁺ currents through inactivated Kv channels support the conclusions made from the fluorescence data obtained from Shaker channels described here. In the Na⁺ experiments there was a large decrease in open-state current at low pH, but large slow inward Na⁺ tail currents were maintained upon repolarization. This supports the idea that channels can still activate and deactivate at low pH without passing through the open state, and thus activation gating is expected to be maintained. This is supported by the gating current measurements in Fig. 2 as well as the fluorescence experiments in Figs. 1 and 3, which imply that low pH does not alter total gating charge movement. As well, these Na⁺ permeation experiments showed an equivalent Na⁺ conductance through open-inactivated channels during sustained depolarizations at normal and acidic pH. These data suggest the possibility that channels are closed-inactivated at low pH, and are able to activate directly into Na⁺- conductive depolarization-induced inactivated states without actually opening. This idea is consistent with the present fluorescence and single channel experiments, which show directly that a population of channels becomes unavailable during acidic pH (Figs. 5, 6, and 10) due to the stabilization of inactivation from resting closed channel states (Figs. 10–13), making them unable to open. A kinetic model to describe the Na⁺ permeation experiments suggested that channels accumulated in the open-inactivated state during depolarization and made the transition to closed-inactivated states via an intermediate "R" state in the recovery pathway (Zhang et al., 2003). Channels could not reopen from resting closed-inactivated states in the model (Zhang et al., 2003), but it is not clear whether or not Na⁺ as the permeant ion causes other structural changes within the selectivity filter that do not allow it to accurately model activation of closed-inactivated channels with K⁺ as the charge carrier (Panyi and Deutsch, 2006).

It has previously been shown that Shaker channels may undergo a type of closed-state inactivation called U-type inactivation (Klemic et al., 2001) and it is helpful to consider whether this is the closed state stabilized at low pH. In U-type inactivation, maximal current reduction occurs at intermediate potentials, where the dwell time in intermediate states is longer, and less inactivation occurs at more depolarized potentials when the open probability is high, resulting in a characteristic U-shaped inactivation–voltage relationship. However, although the fluorescence data strongly support closed-state inactivation, the effects of raising external K⁺ reported here are opposite to those expected for U-type inactivation. U-type inactivation is enhanced by raising external K⁺ (Klemic et al., 2001), whereas we show in Fig. 10 that high external K⁺ rescues channels from closed-state inactivation, which suggests that the pH-induced closed-state inactivation is unlikely to be U-type in nature.

	ACKNOWLEDGMENTS

 
This work was supported by grants from the Heart and Stroke Foundation of British Columbia and Yukon and the Canadian Institutes of Health Research to D. Fedida and S.J. Kehl. D. Fedida is supported by a Career Investigator award from the Heart and Stroke Foundation of British Columbia and Yukon. T.W. Claydon was supported by a postdoctoral research fellowship funded by a Focus on Stroke strategic initiative from The Canadian Stroke Network, the Heart and Stroke Foundation, the CIHR Institute of Circulatory and Respiratory Health, and the CIHR/Rx&D Program along with AstraZeneca Canada. M. Vaid was supported by a Michael Smith Foundation for Health Research Studentship. S. Rezazadeh was supported by a Heart and Stroke Foundation of British Columbia and Yukon Studentship and a University of British Columbia Graduate Fellowship.

Olaf S. Andersen served as editor.

Submitted: 2 March 2007
Accepted: 6 April 2007

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