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NEUROPHARMACOLOGY

4-Aminopyridine Prevents the Conformational Changes Associated with P/C-Type Inactivation in Shaker Channels

Departments of Anesthesiology, Pharmacology & Therapeutics (T.W.C., D.F.) and Cellular and Physiological Sciences (M.V., S.R., S.J.K.), University of British Columbia, Vancouver, British Columbia, Canada

Received for publication July 6, 2006
Accepted September 29, 2006.

	Abstract

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The effect of 4-aminopyridine (4-AP) on K_v channel activation has been extensively investigated, but its interaction with inactivation is less well understood. Voltage-clamp fluorimetry was used to directly monitor the action of 4-AP on conformational changes associated with slow inactivation of Shaker channels. Tetramethylrhodamine-5-maleimide was used to fluorescently label substituted cysteine residues in the S3-S4 linker (A359C) and pore (S424C). Activation- and inactivation-induced changes in fluorophore microenvironment produced fast and slow phases of fluorescence that were modified by 4-AP. In Shaker A359C, 4-AP block reduced the slow-phase contribution from 61 ± 3 to 28 ± 5%, suggesting that binding inhibits the conformational changes associated with slow inactivation and increased the fast phase that reports channel activation from 39 ± 3 to 72 ± 5%. In addition, 4-AP enhanced both fast and slow phases of fluorescence return upon repolarization ( reduced from 87 ± 15 to 40 ± 1 ms and from 739 ± 83 to 291 ± 21 ms, respectively), suggesting that deactivation and recovery from inactivation were enhanced. In addition, the effect of 4-AP on the slow phase of fluorescence was dramatically reduced in channels with either reduced (T449V) or permanent P-type (W434F) inactivation. Interestingly, the slow phase of fluorescence return of W434F channels was enhanced by 4-AP, suggesting that 4-AP prevents the transition to C-type inactivation in these channels. These data directly demonstrate that 4-AP prevents slow inactivation of K_v channels and that 4-AP can bind to P-type-inactivated channels and selectively inhibit the onset of C-type inactivation.

View larger version (39K): [in this window] [in a new window]   Fig. 1. Relevant sites within the Shaker channel and kinetic scheme of the action of 4-AP. Cartoon of the Shaker channel (A) with amino acid positions that are relevant to the present study are highlighted. The native cysteine residue in the S1-S2 linker was replaced with a valine residue (C245V). W434F and T449V were used to modify inactivation. TMRM was attached at A359C in the S3-S4 linker or S424C in the outer pore. The molecular structures of 4-AP and TMRM are also shown. A 4-AP molecule is 2.4 Å wide and 3.6 Å long. The diameter of the inner vestibule of the pore is 12 Å. A TMRM molecule is 14 Å across. Kinetic scheme for the action of 4-AP (B) was modified from that of Armstrong and Loboda (2001). C0 to C4 represent closed states of the channel, and O represents the open state. Transitions between C0 and C4 represent voltage-dependent early independent gating transitions, and the transition from C4 to O (*) represents the weakly voltage-dependent final concerted opening. Hence, C4 can be considered the recently described activated-not-open state (del Camino and Yellen, 2005; Pathak et al., 2005). 4-AP can only bind when channels open, and once bound, the drug strongly biases the closed 4-AP-bound state (C4,AP). 4-AP-bound channels are able to undergo the independent transitions (C0,AP to C4,AP), but the drug retards the transition to opening. Inactivation gating transitions from the open state shown are as suggested previously (Loots and Isacoff, 1998). IP and IC represent P- and C-type-inactivated states, respectively. The dashed arrows reflect that the transitions made during recovery from inactivation are not known. P-type inactivation from the open state only is shown, although there is evidence for P-type inactivation from rest (Yang et al., 1997). The ability of 4-AP to bind to P-type-inactivated channels (IP to IP,AP) is demonstrated in the present study by actions on W434F mutant channels. The fate of IP,AP channels was not directly addressed in the study, so the unbinding transitions from this state have been omitted for reasons of clarity and to avoid undue speculation.

In support of this idea, 4-AP causes very subtle changes in channel on-gating currents (Loboda and Armstrong, 2001) but very profound changes in off-gating (McCormack et al., 1994; Bouchard and Fedida, 1995; Loboda and Armstrong, 2001). Specifically, 4-AP prevents the slowing of charge return that occurs on repolarization from potentials that cause channel opening, which suggests that 4-AP selectively blocks the concerted opening step in the activation sequence without altering the early independent gating transitions (Armstrong and Loboda, 2001; Pathak et al., 2005; del Camino et al., 2005). Further support for this mechanism of action comes from experiments using the Shaker ILT mutation (Pathak et al., 2005), which energetically dissociates the early independent gating transitions from the last concerted opening transition (Smith-Maxwell et al., 1998a,b). Figure 1B shows a gating scheme [simplified from that of Armstrong and Loboda (2001)] that summarizes these actions of 4-AP on activation gating.

Although the disruption of channel activation by 4-AP has been extensively investigated, as described above, the relationship between 4-AP block and channel inactivation is less well understood. Castle et al. (1994) used complex voltage-clamp protocols to demonstrate that inactivation and 4-AP binding to Shaker and K_v1.1 channels were mutually exclusive; however, since then, few additional studies have been performed. The recent development of site-directed voltage-clamp fluorimetry allows the real-time monitoring of conformational changes associated with channel gating that do not result in ionic current (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). This is accomplished by examining the changes in the emission from fluorescently labeled residues, brought about by modifications of the fluorophore microenvironment as the channels activate and inactivate (Loots and Isacoff, 1998; Bezanilla, 2002). By attaching tetramethylrhodamine-5-maleimide (TMRM) at A359C in the S3-S4 linker and at S424C in the outer pore (Fig. 1A), the conformational rearrangements associated with movement of the voltage sensor during channel activation and of the pore during inactivation can be directly observed. For example, fluorescence changes from TMRM attached at A359C occur with the same voltage dependence as movement of the voltage sensor charge (Mannuzzu et al., 1996; Cha and Bezanilla, 1997), and the slow fluorescence deflection from TMRM attached at S424C occurs on a similar timescale to slow inactivation and is sensitive to manipulations that alter its rate (Loots and Isacoff, 1998). Here, we have used this technique to scrutinize directly the effect of 4-AP on slow inactivation of Shaker channels. Relevant amino acid sites used in this study are shown in Fig. 1A. We show that 4-AP prevents the conformational changes in Shaker channels associated with inactivation. Furthermore, we show that 4-AP can bind to P-type-inactivated channels and inhibit the onset of C-type inactivation.

	Materials and Methods

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Solutions and Chemicals
Barth's medium contained 88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO₃, 0.82 mM MgSO₄, 0.33 mM Ca(NO₃)₂, 0.41 mM CaCl₂, and 20 mM HEPES, titrated to pH 7.4 using NaOH. ND96 bath solution contained 96 mM NaCl, 3 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, and 5 mM HEPES, titrated to pH 7.4 using NaOH. TMRM labeling of oocytes was performed in a depolarizing solution that contained 98 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, 5 mM HEPES, titrated to pH 7.4 using KOH, and 50 µM TMRM. Working concentrations of 4-AP were diluted from a 100 mM stock solution made with ND96 solution and titrated to pH 7.4 using NaOH. All chemicals were purchased from Sigma-Aldrich (Mississauga, ON, Canada).

Molecular Biology and RNA Preparation
A modified pBluescript SKII oocyte expression vector (pEXO) was used to express the N-terminal deletion mutant Shaker 6–46 (a kind gift from Dr. A. Sivaprasadarao) that is fast-inactivation-removed (Hoshi et al., 1991), and the pBluescript SKII expression vector was used to express Shaker 6–46 ILT (a kind gift from Dr. E. Isacoff).

The A359C and S424C mutations were made in the background of a mutation that replaced the only externally accessible cysteine residue, found in the S1-S2 linker, with a valine residue (C245V). This was done to reduce nonspecific labeling and to introduce a cysteine residue for site-specific labeling with TMRM in either the S3-S4 linker (A359C) or the outer pore (turret) region (S424C). 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, throughout this article. The T449V mutation in Shaker was used to inhibit slow inactivation (Lopez-Barneo et al., 1993), and the W434F mutation was used to permanently inactivate channels (Perozo et al., 1993; Yang et al., 1997). Point mutations were generated using the Stratagene QuikChange kit (Stratagene, La Jolla, CA). All primers used were synthesized by Sigma-Genosys (Oakville, ON, Canada). All constructs were sequenced using the core facility unit at the University of British Columbia. cDNA was linearized using BstEII (for Shaker 6–46 channels) or HindIII (for Shaker 6–46 ILT channels). cRNA was synthesized from linear cDNA using the T7 mMessage mMachine T7 Ultra cRNA transcription kit (Ambion, Austin, TX).

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) of cRNA using a Drummond digital microdispenser (Fisher Scientific, ON, Canada) and then incubated in Barth's medium at 19°C. Currents were recorded 1 to 7 days after injection.

Voltage-Clamp Fluorimetry
Labeling of introduced cysteine residues was performed with TMRM (Invitrogen, Carlsbad, CA), which reacts specifically with cysteine residues and has a maximal light absorption at 542 nm and a maximal emission at 567 nm. Oocytes were washed and labeled with 50 µM TMRM in oocyte-depolarizing solution for 30 min at 10°C in the dark. Oocytes were then placed in ND96 solution in the dark until they were voltage-clamped at room temperature. Fluorimetry was performed using a Nikon TE300 inverted microscope with EpiFluorescence attachment and a 9124b Electron Tubes photomultiplier tube (PMT) module (Cairn Research, Kent, UK). The TMRM dye was excited by light from a 100-W mercury lamp that was filtered with a 525-nm band pass excitation filter and passed via a dichroic mirror and 20x objective to the oocyte in the bath chamber. Fluorescence emission from the dye was collected via the same 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 pClamp8 software (Axon Instruments, Foster City, CA) to record the fluorescence emission intensity. Fluorescence signals were filtered at 1 kHz with a sampling frequency of 20 (when the pulse duration was 100 ms), 10 (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 majority of bleaching of the fluorescence signal during 7- or 42-s pulses, the fluorescence recorded, during a pulse to –80 mV where there was no voltage-dependent deflection, was subtracted. Simultaneous voltage clamp of the oocyte and acquisition of the current and voltage signals were achieved using the two-microelectrode voltage-clamp technique with a Warner Instruments OC-725C amplifier (Hamden, CT), Axon Digidata 1322, and pClamp8 software. Microelectrodes were filled with 3 M KCl and had a resistance of 0.5 to 1.5 M.

Data Analysis
Conductance-voltage relation (G-V) curves throughout the text were derived using the normalized chord conductance, which was calculated by dividing the maximal current during a depolarizing step by the driving force derived from the K⁺ equilibrium potential (internal [K⁺] was assumed to be 98 mM). G-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.

The Hill equation was used to fit the concentration-response relationship of the effect of 4-AP using Prism 3.02 (GraphPad Software, Inc., San Diego, CA):

(2)

where y is the fraction of current remaining at a given membrane potential, IC₅₀ is the concentration required to achieve half-maximal block, [4-AP] is the concentration of 4-AP in the bath solution, and n_H is the Hill coefficient. Data throughout the text and figures are shown as means ± S.E.M.

	Results

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Shaker A359C TMRM Fluorescence Can Report Both Channel Activation and Inactivation. The slow decay of K_v channel current during prolonged depolarizations involves at least two consecutive steps (see Yellen, 1998; Kurata and Fedida, 2006), the first step closing the outer pore (P-type inactivation) and allowing subsequent stabilization of the inactivated state (C-type inactivation), which is reported by stabilization of the activated conformation of the voltage sensor (DeBiasi et al., 1993; Olcese et al., 1997; Yang et al., 1997; Loots and Isacoff, 1998, 2000) (also see Fig. 1B). Although these processes were first described using standard voltage-clamp techniques, changes of channel conformation during P- and C-type inactivation can also be clearly observed using voltage-clamp fluorimetry (Loots and Isacoff, 1998, 2000). In particular, two sites effectively report the conformational changes occurring during slow inactivation: Ala359 in the S3-S4 linker and Ser424 in the pore. Data in Fig. 2A show typical current and fluorescence recordings from Shaker A359C channels (Fig. 2, A and B) and two Shaker mutants with disrupted inactivation, T449V (Fig. 2, D and E) and W434F (Fig. 2, G and H). Currents are recorded during 100-ms pulses to a range of potentials that activate ionic current; the currents activate rapidly and, at the more positive potentials, decay (inactivate) during the sustained clamp pulse. On the time base studied here, the extent of the decay is small but clearly seen in comparison to the horizontal dotted line that shows a single exponential fit of the rising phase of current. Fluorescence recordings (Fig. 2B) obtained simultaneously with the currents in panel A show a similar rapid onset manifested here as a decrease in fluorescence followed at the more positive clamp potentials by a slower declining phase, which corresponds to the current decay seen in Fig. 2A. It has been suggested that the rapid fluorescence decrease reported by A359C represents channel activation, whereas the slower decay phase reflects channel inactivation (Loots and Isacoff, 1998). The amplitudes of the fast (F_fast) and slow (F_slow) phases of the fluorescence report from Fig. 2B are plotted together with the activating and inactivating components of the ionic current record as a function of the potential during the clamp step in Fig. 2C. It can be seen that the rapid phase of fluorescence change, F_fast, is negatively shifted from the G-V on the voltage axis as it reports the voltage sensor movement that precedes channel opening (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). In Shaker A359C channels, F_fast contributes 64 ± 7% to the total fluorescence deflection during a 100-ms pulse to +100 mV (n = 10). The voltage dependence of the slow phase of fluorescence deflection, F_slow, is displaced to the right of the fast fluorescence relationship and shows a voltage dependence similar to that for the G-V relation and also the inactivating component of current (I_inact), which is a plot of the amplitude of the current that decays during the pulse at each test potential (I_inact at each potential is normalized to that at +100 mV). This is expected because inactivation is coupled to channel opening and should approximate the G-V relation. F_slow contributes 36 ± 7% to the total fluorescence deflection (n = 10). The idea that the slow phase of fluorescence deflection is associated with channel inactivation is supported by experiments on two mutant channels with altered properties of slow inactivation (Fig. 2, D–I).

View larger version (39K): [in this window] [in a new window]   Fig. 2. A fluorophore attached at A359C in the Shaker channel reports both activation and inactivation. Typical current traces (A, D, and G) and fluorescence signals (B, E, and H) recorded from Shaker A359C channels without or with mutations (T449V or W434F) that affect inactivation properties during 100-ms voltage pulses from –100 to +100 mV from a holding potential of –80 mV are shown. Although pulses were applied in 10-mV increments, only every other pulse is shown for clarity. Dashed lines show extrapolations of single exponential fits (from when 50% of the current has activated) to the time course of current activation (A and D) or the fast fluorescence deflection (B, E, and H) at +100 mV to highlight the degree of ionic current and fluorescence decay in each channel. C, F, and I, Ffast and Fslow phases of the fluorescence signals and the activating (G-V) and inactivating (Iinact) components of the ionic current record plotted as a function of potential (n = 4–10). Iinact represents the amplitude of the current that decays during the 100-ms pulse. This value at each potential is normalized to that measured at +100 mV. The fluorescence reports from Shaker T449V A359C and Shaker W434F A359C channels could be fit with a single exponential that represented just the fast phase; therefore, Fslow is absent. Shaker W434F A359C channels were nonconducting; therefore, the G-V relationship is absent in this mutant.

The mutation T449V in the outer mouth of the Shaker channel pore has long been used to disrupt slow inactivation (Lopez-Barneo et al., 1993). Current records in Fig. 2D and the voltage relations in Fig. 2F show that, as expected, the slow phase of current decay attributable to slow inactivation is largely prevented in this channel (i.e., I_inact is absent on this timescale). Consistent with this, the slow phase of fluorescence decay recorded from A359C is also absent (Fig. 2, E and F). A second mutation, W434F, is known to induce permanent P-type inactivation (Perozo et al., 1993; Yang et al., 1997; Olcese et al., 1997; Loots and Isacoff, 1998), and there are no ionic currents recordable from this mutant (Fig. 2G); however, the fluorescence report is still robust (Fig. 2H) as activation proceeds normally despite the channels being inactivated (Perozo et al., 1993; Olcese et al., 1997; Starkus et al., 1998). It can be seen that, although the fast phase of fluorescence is preserved, the slow fluorescence changes are completely abolished (Fig. 2, H and I). Taken together, the results in Fig. 1 clearly demonstrate that the fluorescence report from A359C can track conformational changes associated with Shaker channel activation and, of particular importance for the purposes of this study, channel inactivation. Even though channels may be unable to conduct, either before opening or after inactivation, the fluorophore remains able to report changes in channel gating state.

The Effect of 4-AP on Channel Inactivation. Using voltage-clamp fluorimetry, we investigated the effect of 4-AP on the conformational rearrangements associated with channel inactivation by recording ionic currents and fluorescence signals from TMRM attached at A359C during 7-s depolarizing pulses to +60 mV in the absence and presence of 3 mM 4-AP (Fig. 3A). In the control trace, channels open rapidly upon depolarization; however, during maintained depolarization, the current declines due to the onset of slow (P- and then C-type) inactivation. The signal from the fluorophore in the S3-S4 linker (Fig. 3A, bottom) reports on the conformational changes associated with these gating events. On depolarization, there is a rapid fluorescence deflection followed by a slowly declining phase, which displays kinetics similar to the decay of ionic current and represents the transition of channels into the inactivated conformation as described in Fig. 2. This is highlighted in Fig. 3B, which shows superimposed traces of ionic current and fluorescence signal decay recorded simultaneously from the same cell. The mean time constants of decay for the ionic current and fluorescence signal from 14 such experiments are very similar; = 2.7 ± 0.2 and = 1.9 ± 0.2 s, respectively. On repolarization to the holding potential, the ionic currents rapidly deactivate and channels recover from inactivation. Again, these gating events are reported by the fluorophore (Fig. 3C). On repolarization, the fluorescence returns toward baseline and shows a fast phase that has a of 87 ± 15 ms and reflects the time course of the return of the voltage sensor during channel deactivation and a slow phase that has a of 739 ± 82 ms, which reflects the slower return of the voltage sensor to its resting state following stabilization of its activated conformation during inactivation (Cha and Bezanilla, 1997; Loots and Isacoff, 1998).

View larger version (19K): [in this window] [in a new window]   Fig. 3. 4-AP abolishes the inactivation-dependent component of fluorescence. A and D, ionic currents (top) and fluorescence signals (bottom) recorded during a 7- (A) or 42-s (D) depolarizing pulse to +60 mV from Shaker A359C (A) or Shaker S424C (D) channels from a holding potential of –80 mV in the absence and presence of 4-AP. The dotted trace represents the normalized ionic current in the presence of 4-AP scaled 2-fold in A and 8-fold in D. B and E, superimposition of ionic current and the slow phase of the fluorescence deflection in the absence of 4-AP show similar kinetics. The time constants of ionic current and fluorescence signal decay are 2.5 ± 0.2 and 1.9 ± 0.2 s, respectively (n = 16), for Shaker A359C in B and 2.6 ± 0.3 and 3.8 ± 0.9 s, respectively (n = 6–10), for Shaker S424C in E. C and F, normalization of the return of the fluorescence signal to baseline on repolarization in the absence and presence of 4-AP from Shaker A359C (C) and Shaker S424C (F). The return of the fluorescence is biexponential. The fast and slow time constants are 87 ± 15 (relative amplitude, a = 0.64 ± 0.07) and 739 ± 82 ms (a = 0.36 ± 0.07), respectively, in the absence of 4-AP and 40 ± 1(a = 0.75 ± 0.04) and 291 ± 21 ms (a = 0.25 ± 0.04), respectively, in the presence of 3 mM 4-AP in Shaker A359C. The corresponding values in Shaker S424C are 269 ± 40 ms (a = 0.39 ± 0.10) and 5.2 ± 1.1 s (a = 0.61 ± 0.10) in the absence of 4-AP and 181 ± 23 ms (a = 0.45 ± 0.10) and 6.1 ± 1.0 s (a = 0.55 ± 0.10) in the presence of 10 mM 4-AP.

In the presence of 4-AP, there were marked changes in both the ionic currents and fluorophore reports of both A359C and S424C TMRM-labeled mutant channels. The ionic currents are partially blocked, and the rate of the decay is apparently reduced. Peak current is inhibited by 65 ± 6% at 3 mM 4-AP (n = 5). Scaling of the ionic traces in the presence of 4-AP to the control trace in each mutant (Fig. 3, A and D, dotted lines) shows that the rate and extent of slow inactivation of ionic current are significantly reduced in the presence of 4-AP. Fluorophore signals during prolonged depolarization recorded in the presence of 4-AP from both channels (Fig. 3, A and D) show, for the first time, the conformational changes associated with the inactivation of drug-bound channels. 4-AP increases the relative contribution of the fast phase of fluorescence change in Shaker A359C channels from 39 ± 3 to 72 ± 5% and reduces the contribution of slow secondary fluorescence deflection that was associated with channel inactivation from 61 ± 3 to 28 ± 5% (n = 5; paired t test, P < 0.001). This results in a crossover of the fluorescence traces because 4-AP increases the instantaneous fluorescence deflection by 54 ± 29% but reduces the deflection at the end of the depolarizing pulse by 28 ± 9% (Fig. 3A, bottom). In addition, the fast and slow phases of the return of the fluorescence to baseline on repolarization are markedly accelerated in the presence of 4-AP (Fig. 3C). The fast is reduced to 40 ± 1 ms (from 87 ± 15 ms without 4-AP), suggesting that deactivation is enhanced, which is consistent with previous observations that 4-AP abolishes the immobilization of charge return (McCormack et al., 1994; Bouchard and Fedida, 1995; Loboda and Armstrong, 2001), and the slow is reduced to 291 ± 21 ms (from 739 ± 82 ms without 4-AP), which is consistent with the view proposed by Castle et al. (1994) that fewer channels inactivate in the presence of 4-AP during the maintained depolarization. In the case of the Shaker S424C channel, the fluorescence signal reporting inactivation is reduced by 64 ± 5% (n = 4) in the presence of 4-AP (Fig. 3D, bottom). An increase in the fast phase is not observed with Shaker S424C, probably because the fluorophore at this position does not report as well on activation (Loots and Isacoff, 1998). Interestingly, the time constants of the fast and slow phases of the return of the fluorescence signal upon repolarization are not different from those in the absence of 4-AP (Fig. 3F). The of the fast phase is 181 ± 23 ms, and that of the slow phase is 6.1 ± 1.0 s (n = 5). Taken together, these current and fluorescence data, which allow direct real-time observation of the conformational changes associated with inactivation, confirm that channel inactivation is greatly reduced after 4-AP binding.

To examine the interdependence of 4-AP block and P/C-type channel inactivation further, the effect of 4-AP binding to T449V and W434F channels was examined. Data in Fig. 4A show ionic currents and fluorescence signals recorded during 7-s depolarizing pulses to +60 mV from a holding potential of –80 mV. As suggested earlier (Fig. 2), the T449V mutation largely prevents inactivation, although some current decay does still persist during a long depolarizing pulse ( = 4.4 ± 0.7 s and the relative amplitude of the decaying component, a = 0.22 ± 0.03 in Shaker T449V A359C compared with = 2.5 ± 0.2 s and a = 0.69 ± 0.03 in Shaker A359C; n = 5; P < 0.01, t test). The reduced inactivation of T449V mutant channels is accompanied by a slowed and reduced fluorescence decay ( = 2.4 ± 0.3 s compared with = 1.9 ± 0.2 s without the mutation; n = 5–16). Similar to Shaker A359C channels, 3 mM 4-AP blocked Shaker T449V A359C ionic current by 76 ± 2% (Fig. 4A). However, the fluorophore report shows that 4-AP has a much smaller effect on both the fast and slow phases of fluorescence deflection. The extent of the divergence of the fluorescence traces is reduced, because 4-AP increases the instantaneous fluorescence deflection only 21 ± 7% (compared with 54 ± 29% in the absence of 4-AP) and reduces the deflection at the end of the pulse by only 13 ± 12% (compared with 28 ± 9% in the absence of 4-AP). These data suggest that the reduced inactivation of the T449V mutant diminishes the scope for the effect of 4-AP on the fluorescence signals. The reduction of the small slow fluorescence deflection from Shaker T449V A359C in the presence of 4-AP (Fig. 4A) is probably due to inhibition of the residual inactivation present in these mutant channels. The fast phase of the fluorescence return of Shaker T449V A359C channels is still accelerated by 4-AP from 43 ± 9 to 9 ± 1 ms (n = 3) because 4-AP enhances deactivation, and the of the slow phase of the fluorescence return is also reduced from 775 ± 132 to 377 ± 56 ms (n = 3). These data show that 4-AP can block currents from inactivation-reduced T449V channels equivalently to A359C channels and that 4-AP can still inhibit the residual inactivation present in T449V channels, although the fluorescence report is much less altered by 4-AP in these channels.

View larger version (15K): [in this window] [in a new window]   Fig. 4. Modulation of inactivation alters the effect of 4-AP on Shaker A359C TMRM fluorescence. A and C, ionic (top) and fluorescence (bottom) traces recorded from Shaker T449V A359C inactivation-reduced mutant channels (A) and Shaker W434F A359C permanently inactivated channels (C) during a 7-s depolarizing pulse to +60 mV from a holding potential of –80 mV in the absence and presence of 4-AP. The dotted trace in A represents the ionic current in the presence of 4-AP scaled 4-fold to match the peak of the control current. B and D, normalization of the return of the fluorescence signal on repolarization in the absence and presence of 4-AP from Shaker T449V A359C (B) and Shaker W434F A359C (D). The fast and slow time constants are 43 ± 9 (relative amplitude, a = 0.71 ± 0.04) and 775 ± 132 ms (a = 0.29 ± 0.04), respectively, in the absence of 4-AP and 9 ± 1 (a = 0.88 ± 0.05) and 337 ± 56 ms (a = 0.12 ± 0.05), respectively, in the presence of 3 mM 4-AP in Shaker T449V A359C. The corresponding values in Shaker W434F A359C are 49 ± 15 (a = 0.68 ± 0.05) and 474 ± 3 ms (a = 0.32 ± 0.05) in the absence of 4-AP and 23 ± 1 (a = 0.86 ± 0.06) and 167 ± 65 ms (a = 0.14 ± 0.06) in the presence of 3 mM 4-AP.

In permanently inactivated W434F channels, ionic currents cannot be recorded (Figs. 2G and 4C). Despite this, depolarization produces rapid quenching of the large TMRM fluorescence signal from A359C. As described in Fig. 2, the fluorophore reports very little secondary slow phase, and Fig. 4C shows that this is the case even during 7-s depolarizing pulses. The contribution of the fast phase to the total fluorescence is 95 ± 2% (Fig. 4C). The application of 4-AP has no effect on the kinetics of fluorophore movement during the depolarization, except for a small reduction in the absolute signal. In the presence of 3 mM 4-AP, the fast phase contributes 90 ± 4% to the total deflection (n = 3), which is not different from that without 4-AP. As in Shaker A359C channels (Fig. 3C), 4-AP accelerates the fast phase of the return of the fluorescence signal on repolarization (Fig. 4D). The of the fast phase is reduced from 49 ± 15 ms in the absence to 23 ± 1 ms in the presence of 3 mM 4-AP (n = 3), which is consistent with the previous observation that 4-AP abolishes immobilization of the voltage sensor of W434F channels on repolarization (McCormack et al., 1994). Although there is no slow secondary fluorescence deflection from W434F channels during a 7-s depolarization, the recovery of the fluorescence signal on repolarization shows a slow phase with a of 474 ± 3 ms that probably represents the recovery of channels that become C-type-inactivated during the voltage pulse. This is consistent with previous observations that P-type-inactivated W434F channels are able to undergo C-type inactivation during prolonged depolarization (Olcese et al., 1997). Interestingly, 4-AP accelerates the slow phase of fluorescence return on repolarization (Fig. 4D). The of the slow phase of fluorescence return is reduced to 167 ± 65 ms in the presence of 3 mM 4-AP (n = 3). The incomplete return of the fluorescence signal to the baseline in this case may reflect minor bleaching of the signal rather than incomplete voltage sensor return, and this is supported by the evidence of similar incomplete fluorescence return observed in other previous records (e.g., Loots and Isacoff, 1998). These data suggest that 4-AP may be able to bind to P-type-inactivated channels and inhibit, at least in part, the transition to C-type inactivation.

The Onset of 4-AP Block Detected by Changes in Shaker A359C TMRM Fluorescence. As described in Fig. 3A, 4-AP introduces a crossover of the fluorescence signals from Shaker A359C channels in the early stages of a maintained depolarization. To investigate this phenomenon further, we measured the effect of different concentrations of 4-AP on the ionic current and fluorescence report during 200-ms voltage pulses to +60 mV (Fig. 5A). In each case in Fig. 5A, the control trace in the absence of 4-AP is shown along with the 1st and 5th pulses (at 0.2 Hz) after a 3-min application of the indicated concentration of 4-AP, during which channels were held closed (holding potential –80 mV). It has been shown that block cannot occur until a channel is opened for the first time (1st pulse) in the presence of the drug (Choquet and Korn, 1992; McCormack et al., 1994; Bouchard and Fedida, 1995). Steady-state block is reached by the 5th pulse in each case. In the presence of 0.3 mM 4-AP, the peak ionic current amplitude on the 1st pulse is not different from that of control; however, there is a slow decline in the current during the pulse that represents the onset of channel block (Fig. 5A, left). The fluorescence signals in Fig. 5A accurately report the gating events associated with this onset of channel block. The instantaneous fluorescence signal recorded on the 1st pulse in the presence of 0.3 mM 4-AP (Fig. 5A, right) is identical to that in control conditions, both in terms of amplitude and kinetics, and is followed by a clear decrease of fluorescence that coincides with the decline of ionic current and saturates once steady-state block is reached by the 5th pulse (Fig. 5A, right). Increasing concentrations of 4-AP exaggerate both the decay of the ionic current and the increase of the fluorescence signal (Fig. 5A). Because 4-AP does not significantly quench TMRM directly (see Discussion), these data suggest that binding of 4-AP directly increases the fluorescence signal from Shaker A359C channels during short depolarizations by inhibiting ongoing inactivation in channels that normally occurs once they open. This allows the fast phase of fluorescence from opening channels to be more completely detected. Consistent with this, in a different set of experiments measuring the onset of block during 7-s pulses, the contribution of the fast-phase increases and slow-phase decreases by 3 ± 1% with 0.3 mM 4-AP, 4 ± 3% with 1 mM, 10 ± 2% with 3 mM, and 14 ± 1% with 10 mM (n = 3–4; when compared with the amplitudes of the fast and slow phases in the absence of 4-AP).

View larger version (43K): [in this window] [in a new window]   Fig. 5. The onset of 4-AP block detected by changes in Shaker A359C TMRM fluorescence. A, ionic currents (left) and fluorescence signals (right) recorded during depolarizing pulses to +60 mV from Shaker A359C in the absence of 4-AP (control) and the 1st and 5th pulses following a 3-min application of 4-AP (the membrane was held at –80 mV during 4-AP application to prevent channel block). B, 4-AP concentration-response curves generated from the steady-state level of block using ionic currents or fluorescence deflections at +60 mV as shown in A. The IC50 and nH calculated from the ionic currents are 1.3 ± 0.1 and 0.92 ± 0.05 mM, respectively (n = 3–6). C, G-V relations with increasing concentrations of 4-AP were constructed using the peak current amplitudes during the pulse (n = 3–5). V1/2 and k values were –10.0 ± 1.8 and 20.5 ± 1.5 mV for control, 3.3 ± 2.6 and 26.3 ± 2.0 mV for 300 µM 4-AP, 9.7 ± 2.4 and 23.9 ± 1.8 mV for 1 mM 4-AP, 37.9 ± 3.6 and 29.0 ± 2.0 mV for 3 mM 4-AP, and 49.3 ± 4.5 and 29.1 ± 2.2 mV for 10 mM 4-AP. These values in the absence of 4-AP are similar to those previously reported for channels with mutations of the S3-S4 linker (Gonzalez et al., 2001) D, F-V relations were generated by normalizing the peak fluorescence deflections in a given 4-AP concentration to those in the absence of 4-AP in each oocyte (n = 3–5). The V1/2 and k values were –26.7 ± 1.5 and 26.2 ± 1.3 mV, respectively, for control, –35.5 ± 1.7 and 23.2 ± 1.4 mV for 300 µM 4-AP, –39.0 ± 1.1 and 19.7 ± 0.9 mV for 1 mM 4-AP, –38.0 ± 1.3 and 18.0 ± 1.1 mV for 3 mM 4-AP, and –26.9 ± 1.8 and 24.2 ± 1.5 mV for 10 mM 4-AP.

Concentration-response curves calculated from steady-state currents at +60 mV, such as those shown in Fig. 5A, are shown in Fig. 5B. 4-AP inhibits ionic current with an IC₅₀ of 1.3 ± 0.1 mM and an n_H of 0.92 ± 0.05 (n = 3–6). This is similar to our own measurements as well as previous reports of 4-AP block of Shaker channels that lack an engineered cysteine residue and TMRM dye molecule in Xenopus oocytes, which suggest an IC₅₀ of 0.1 to 0.3 mM (McCormack et al., 1994; Castle et al., 1994) but suggests that the A359C mutation and its attached TMRM molecule may alter 4-AP potency to a small degree. The effect of different concentrations of 4-AP on the steady-state conductance- and fluorescence-voltage relationships is shown in Fig. 5, C and D. These data are taken from currents and fluorescence reports recorded during 200-ms voltage pulses from –100 to +80 mV. It is clear that increasing the concentration of 4-AP shifts the conductance-voltage relation of Shaker A359C channels to more depolarized potentials and also reduces the maximal conductance (Fig. 5C). This is consistent with the previous suggestion that 4-AP binding stabilizes the activated-not-open state of the channel (McCormack et al., 1994; del Camino et al., 2005) where the voltage sensor charges have moved but the concerted opening step is prevented (Pathak et al., 2005). In agreement with this, Fig. 5D shows that 4-AP has no effect on the V_1/2 of the fluorescence-voltage relationship, suggesting that 4-AP has little effect on voltage sensor movement and that 4-AP bound channels gate normally without significant opening, as suggested previously (McCormack et al., 1994; Armstrong and Loboda, 2001). The fluorescence-voltage relationships also show the concentration-dependent increase in the absolute fluorescence signal amplitude seen in the presence of 4-AP in Figs. 3A and 5A without any change in the voltage-dependence of the signal. This suggests that the additional fluorescence signal does indeed reflect activation of channels.

4-AP Stabilizes the Activated-Not-Open State. The ILT mutant Shaker channel provides a useful tool in channel gating studies, because it isolates voltage sensor movement from the cooperative opening transition of channels, i.e., a pulse to 0 mV, for example, will move most of the gating charge but does not culminate in channel opening (Smith-Maxwell et al., 1998a,b) (also see Fig. 1B). This state was recently described as the activated-not-open state (del Camino et al., 2005) and is suggested to be the same state that is stabilized by the binding of 4-AP. We reasoned that the fluorophore report from the ILT mutant channel at 0 mV should therefore be identical to that from 4-AP-bound channels and also insensitive to 4-AP. Figure 6A shows current traces and fluorescence signals recorded from Shaker ILT A359C channels during a 7-s pulse to 0 mV from a holding potential of –80 mV in the absence and presence of 3 mM 4-AP. Figure 6B shows G-V and fluorescence-voltage (F-V) curves plotted from 100-ms pulses over a wide range of potentials. It is clear that at 0 mV the ILT mutant channels do not open because there is no observable current but that the majority of the charge has moved because the fluorophore reports a rapid and large conformational change. Figure 6B shows that the V_1/2 potentials of the G-V and F-V curves of the Shaker ILT A359C channel are separated by more than 150 mV and that opening occurs only at very depolarized potentials. The fluorescence report at 0 mV in the absence of 4-AP shows no secondary movement following the conformational changes associated with charge movement (Fig. 6A) as it does in the Shaker A359C channel in the presence of 4-AP (Fig. 3A). Furthermore, 4-AP does not alter the fluorescence report at 0 mV (Fig. 6A). The similarity between the fluorophore reports from 4-AP bound channels and the ILT mutant channels is consistent with the idea that 4-AP, like the ILT mutant, stabilizes the activated-not-open state of the channel. The lack of evidence of an inactivation transition in the fluorescence trace at 0 mV in the absence of 4-AP also supports the argument that inactivation is strictly coupled to opening in this channel.

View larger version (20K): [in this window] [in a new window]   Fig. 6. 4-AP binding stabilizes the activated-not-open state of the channel. A, ionic (top) and fluorescence (bottom) traces recorded from Shaker ILT A359C channels in the absence and presence of 3 mM 4-AP during a 7-s depolarizing pulse to 0 mV (where maximal voltage sensor movement takes places without any channel opening) from a holding potential of –80 mV. B, isochronal conductance- and fluorescence-voltage relations generated from normalized peak current and fluorescence deflections plotted as a function membrane potential (n = 5). V1/2 and k values were 101.1 ± 5.4 and 36.8 ± 3.5 mV, respectively, for ionic current, and the respective values for the fluorescence deflection were –74.5 ± 0.5 and 16.1 ± 0.4 mV.

	Discussion

Top Abstract Materials and Methods Results Discussion References

 
4-AP Bound Channels Cannot Inactivate. The fluorescence emission of TMRM attached to A359C in the S3-S4 linker reports a fast decrease followed by a slower declining phase on depolarization that reflects activation and inactivation gating conformational changes, respectively (Fig. 2). It is clear that the report from this position is very sensitive to inactivation gating because a slow fluorescence decay can be observed not only during prolonged depolarization (Fig. 3A) when the majority of channels inactivate but also during short pulses (Fig. 2B) when only a small proportion of channels enter the inactivated conformation. Therefore, we used the fluorescence report from Shaker A359C channels to investigate the effect of 4-AP on channel inactivation because, although the effect of 4-AP binding on voltage sensor movement has been extensively studied (McCormack et al., 1994; Bouchard and Fedida, 1995; Armstrong and Loboda, 2001), there is only one report of the effect on inactivation of ionic current, which suggested that inactivation and 4-AP binding are mutually exclusive (Castle et al., 1994). Using the secondary slow phase of fluorescence deflection from Shaker A359C, as well as the slow report from Shaker S424C channels, we have shown directly that 4-AP inhibits the conformational changes associated with inactivation gating of Shaker channels (Fig. 3). In support of this conclusion, the effect of 4-AP on the slow secondary fluorescence deflection that reflects inactivation was reduced in mutant channels with reduced (T449V; Fig. 4A) or permanent (W434F; Fig. 4C) inactivation.

Although it is possible that changes in the fluorescence on 4-AP binding are due to inhibition of ion conduction and that K⁺ permeation alters the fluorescence report, evidence suggests that this is not the case. 1) On repolarization, ionic current reverses direction (as seen in Fig. 3A), but the fluorescence report does not, as would be expected if ion conduction was a major determinant of the fluorescence signal. 2) Previously, Loots and Isacoff (1998) showed that the rate of the slow fluorescence report is highly dependent on modifications that alter the rate of inactivation of ionic current; for example, low pH enhances inactivation and the fluorescence decay, whereas high K⁺ slows inactivation and the fluorescence decay. Consistent with this, the T449V mutation used in the present study (Fig. 4A) slows ionic inactivation and reduces the slow fluorescence decay. Taken together, these data suggest that the fluorescence reports in the present study faithfully report activation and inactivation and are not altered by ion permeation. A similar conclusion was reached by Cha and Bezanilla (1997) who demonstrated that TEA and agitoxin block of the pore altered the fluorescence report from A359C but that this was not due to inhibition of ion conduction, rather that the blocking particles prevented conformational changes associated with channel opening.

4-AP Can Bind to P-Type-Inactivated Channels. Using conventional electrophysiological approaches, Castle et al. (1994) showed that 4-AP slows inactivation of Shaker channels and suggested that 4-AP cannot bind inactivated channels. However, it was not possible in these experiments to separate the effects of 4-AP on P-type inactivation from those on C-type inactivation. In the present study, we have directly observed the effect of 4-AP on P-type inactivation by using the W434F permanently P-type-inactivated mutant channel as well as the effect of 4-AP on C-type inactivation by analyzing the return of the fluorescence signal to baseline upon repolarization, which represents the extent of stabilization of the voltage sensor in its activated conformation, a hallmark of C-type inactivation. W434F channels display almost identical gating currents to wild-type channels but do not conduct K⁺ due to permanent P-type inactivation (Yang et al., 1997). Despite this, channels are thought to be able to access the C-type-inactivated state on depolarization, because they still exhibit charge immobilization following maintained depolarization (Olcese et al., 1997). In the present study, we observed that the return of the fluorescence signal to baseline on repolarization was enhanced in the presence of 4-AP (Fig. 4D). These data suggest that 4-AP can bind to P-type-inactivated channels and inhibit their progression to the C-type-inactivated state (see Fig. 1B). Because 4-AP can only access its binding site in the Shaker channel when the intracellular gate is open, these data show that, although W434F channels are permanently inactivated, the intracellular gate opens with similar voltage dependence to that in conducting channels, which is consistent with the previous observation that the tetraethylammonium binding site is also available on depolarization (Perozo et al., 1993). This conclusion is also consistent with a number of previous studies in which 4-AP is clearly seen to modify gating currents in W434F mutant channels (e.g., McCormack et al., 1994; Bouchard and Fedida, 1995; Loboda and Armstrong, 2001). From our data, we cannot determine whether 4-AP can bind to C-type-inactivated channels. If not, this may be consistent with the inability of 4-AP to bind inactivated channels following the very long pulses used by Castle et al. (1994) that presumably drive most channels to the C-type-inactivated state.

Inactivation Results in Underestimation of Voltage Sensor Movement. When comparing the fluorescence report from Shaker A359C in the absence and presence of 4-AP, we observed a prominent crossover early on in the depolarizing pulse (Fig. 3A). This was reflected as an enhancement of the fluorescence signal during 200-ms pulses (Fig. 5A), suggesting that 4-AP block resulted in an increased quenching of TMRM during depolarization. However, a number of observations suggest that it is unlikely that the increased amplitude of deflection is due to a nonspecific quenching of the TMRM fluorophore by the drug. 1) The basal level of fluorescence emission was the same in the presence and absence of 4-AP; i.e., the increase in the fluorescence signal amplitude only occurs once the channel is opened and 4-AP is allowed to bind. 2) The addition of 4-AP to oocytes expressing channels with no accessible external cysteine residue (i.e., Shaker 6–46 C245V channels) had no effect on fluorophore emission (data not shown). 3) Emission intensities recorded on excitation of TMRM dye in solution with or without 3 mM 4-AP were not different (data not shown). Instead, we conclude that the increased fluorescence deflection seen during the short pulses in Fig. 5A and the crossover of the fluorescence traces in Fig. 3A is a consequence of the 4-AP-induced inhibition of inactivation. The peak of the fluorescence trace in the absence of 4-AP is underestimated due to the onset of inactivation, which results in a slow secondary phase of fluorescence. Because 4-AP binding greatly reduces the ability of Shaker channels to inactivate, this slow phase is almost abolished resulting in an increase in the amplitude of the fast phase of fluorescence. In support of this conclusion, the slow phase of the fluorescence decay in the presence of 4-AP, and the divergence of the fluorescence trace from that in control conditions, is reduced in the inactivation-deficient T449V mutant (Fig. 4A).

4-AP Stabilizes the Activated-Not-Open State. It is well established from gating current studies in Shaker channels that 4-AP binding does not alter the early independent gating transitions between closed states (McCormack et al., 1994; Armstrong and Loboda, 2001). Consistent with this, we have shown that the fast fluorescence report from fluorophores attached in the S3-S4 linker at position A359C is largely unaltered by the binding of 4-AP (Figs. 2B and 5D). It was recently suggested that, like the ILT mutation, 4-AP promotes the activated-not-open state of the channel (Armstrong and Loboda, 2001; del Camino et al., 2005). In the present study, the fluorophore report from ILT mutant channels recorded during a maintained depolarization to 0 mV, which activates channels and moves the voltage sensor (Smith-Maxwell et al., 1998a,b) but does not open channels, mimicked that recorded from normally activating channels in the presence of 4-AP (compare Figs. 3A and 6A). Furthermore, application of 4-AP did not alter this report (Fig. 6). These data support the conclusion that 4-AP binding promotes the activated-not-open channel conformation (del Camino et al., 2005).

	Footnotes

 
This work was supported by grants from the Heart and Stroke Foundations of British Columbia and Yukon and the CIHR (to D.F. and S.J.K.). S.R. was supported by a University of British Columbia Graduate Fellowship. T.W.C 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 Canadian Institute of Health Research (CIHR) Institute of Circulatory and Respiratory Health, and the CIHR/Rx&D Program along with AstraZeneca Canada.

T.W.C. and M.V. contributed equally to this work.

Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.

doi:10.1124/jpet.106.110411.

ABBREVIATIONS: 4-AP, 4-aminopyridine; G, chord conductance; K_v, voltage-gated potassium channels; V_1/2, half-activation potential; F-V, fluorescence-voltage relation; G-V, conductance-voltage relation; PMT, photomultiplier tube; TMRM, tetramethylrhodamine-5-maleimide; ILT, amino acid mutations V369I, I372L, and S376T.

Address correspondence to: Dr. David Fedida, Department of Anesthesiology, Pharmacology and Therapeutics and Cellular and Physiological Sciences, 2350 Health Sciences Mall, Vancouver, BC V6T 1Z3, Canada. E-mail: fedida{at}interchange.ubc.ca

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