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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS
Department of Physiology and Biophysics, School of Medicine and Biomedical Sciences, University at Buffalo, State University of New York, Buffalo, New York
Submitted 18 April 2007 ; accepted in final form 12 June 2007
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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voltage-sensitive potassium channel; Shaker; closed-state inactivation
Due to their important functional roles in the nervous and cardiovascular systems, considerable experimental effort has been exerted to determine the molecular and biophysical basis of Kv4 channel gating transitions. In contrast to Shaker channels, for which the mechanisms of inactivation and recovery from inactivation have been well characterized by an NH2-terminal inactivation ball/peptide domain [N-type inactivation] and external pore vestibule closure [P/C-type inactivation] (1, 5, 24, 25, 31), the mechanisms underlying inactivation and recovery from inactivation in Kv4 channels remain unresolved. Although previous studies (3, 10, 11, 15, 16, 21) have suggested the involvement of both NH2- and COOH-terminal domains, experimental maneuvers used to identify classic N- and P/C-type mechanisms in Kv4 channels have given results either inconsistent with or opposed to those predicted for Shaker channels. For example, increasing extracellular K+ concentration ([K+]o) accelerates inactivation and slows recovery in Kv4.1 [effects opposite to those predicted by the conventional C-type inactivation model (15)], and Kv4.2 NH2-terminal deletion does not remove the fast component of inactivation or alter recovery kinetics (3, 32). In addition, Shaker channels display rapid open-state inactivation with no inactivation from preactivated closed states (1, 31), whereas Kv4 channels can undergo significant closed-state inactivation (CSI) (4, 8, 16, 21, 22, 29).
Additional key differences between Kv1 and Kv4 channels have been noted. With regard to Kv4.3 channels, coexpression of the regulatory 
-subunit KChIP2b significantly accelerated the kinetics of deactivation and recovery from inactivation (22). No such regulation by KChIP isoforms was noted on Kv1.4 channels (17, 19). Also, raising [K+]o from 2 to 98 mM accelerated activation and slowed deactivation kinetics in Kv4.3 channels (30), effects not observed in Shaker channels (13). Taken together, these results raise the possibility that in Kv4.3 channels, voltage-dependent "forward" (activation) and "backward" (deactivation) transitions of the voltage-sensing transmembrane segment S4 may provide a mechanism for not only coupling inactivation to activation but also for coupling recovery to deactivation (21, 22, 23, 29, 30). While such coupling mechanisms have been hypothesized previously (7, 8, 21, 29, 30), their presence in any Kv4 channel has yet to be experimentally verified.
Voltage sensitivity in Shaker channels is conferred to the mechanical work of opening and closing the channel pore primarily by positively charged residues localized to S4 of each of the four identical 
-subunits that comprise the channel (18, 27, 28). This same general mechanism has been assumed to underlie activation in Kv4 channels. However, the steepness of the steady-state activation curves in Kv4 channels is typically two to three times less than those of Shaker channels (16, 21, 22, 29). This is likely due to differences in the number of charged residues within S4: the Kv1.2 channel has six positive charges, whereas the Kv4.3 channel has five. Also, the NH2-terminal region of the S4–S5 linker contains a positively charged lysine in Kv1.2 channels (K312), which is replaced by a glutamine in Kv4.3 channels (Fig. 1A). We thus hypothesized that these differences could account, at least in part, for the variation in basic gating characteristics noted between Shaker and Kv4 channels.
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MATERIALS AND METHODS |
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In vitro transcription and oocyte preparation. Kv4.3 wild-type (WT) and mutant clone plasmids were linearized with the restriction endonuclease XhoI (Promega, Madison, WI). cRNA was synthesized by the mMessage mMachine T7 Ultra Kit (Ambion, Austin, TX). cRNA quantity and quality were evaluated by spectroscopy and agarose gel electrophoresis.
All animal protocols were conducted according to National Institutes of Health-approved guidelines of the Institutional Animal Care and Use Committee of the University at Buffalo, State University of New York. Oocytes were obtained from mature female Xenopus laevis anesthetized by being soaked in 1.0 g/l ethyl-3-aminobenzoate methanesulfonate salt and defolliculated as previously described (22). Twelve to twenty-four hours after isolation, oocytes were injected with 4–9 ng cRNA (Nanoject II, Drummond Scientific, Broomall, PA). Injected oocytes were then incubated for 2–4 days at 18°C.
Electrophysiology. Two-microelectrode voltage-clamp recordings (GeneClamp 500B, Axon Instruments, Union City, CA) were performed on injected oocytes as previously described (22). Recordings (22 ± 2°C) were conducted in ND96 solution [containing (in mM) 96 NaCl, 2 KCl, 1 MgCl2, 1.8 CaCl2, and 5 HEPES; pH 7.40]. All voltage-clamp recordings were conducted at the maximal gain of the amplifier (10,000x) and clamp rise time stability settings of 60–120 µs. Currents were acquired (filtered at 1 kHz and digitized at 5 kHz) with a Digidata 1320A 16-bit acquisition system under pCLAMP 9 software control (Axon Instruments).
Protocols and analysis.
Quantitative analysis was conducted using pCLAMP 9 and Origin 7.5 (OriginLab, Northampton, MA). Analysis of steady-state gating (a4) and iochronal inactivation (i) relationships and kinetic properties (activation, deactivation, inactivation, recovery from inactivation, CSI, and recovery from CSI) were conducted using standard voltage-clamp protocols as previously described (22). The 1-s isochronal inactivation relationship i was determined using a P2 pulse applied to +50 mV preceded by 1-s P1 pulses to varying potentials (see RESULTS for further details). Mean normalized P2 peak currents were fit to the following single Boltzmann relationship: i = 1/{1 + exp[(V – V1/2)/k]}, where V is the test potential (in mV), V1/2 is the potential (in mV) where i = 0.5, and k is the corresponding slope factor (in mV). Estimates of the steady-state activation relationship a4 were generated using a standard saturating tail current protocol (15-ms P1 depolarizing pulses ranging from –95 to +50 mV) followed by a 500-ms P2 pulse applied to –50 mV. In the specific case of R290A, P2 tail currents were measured at either –50 or –120 mV, and the results from both potentials were pooled to construct the mean a4 curve. Normalized mean peak P2 tail currents were fit to the following fourth-order Boltzmann relationship: a4 = 1/{1 + exp[(V – V1/2)/k]}4. This formulation assumes independent activation gating of the four -subunits. Mean values of V1/2 and k for both i and a4 relationships for each expression condition were derived from best-fit analysis using Origin 7.5. Recovery from CSI (see Fig. 8, B and C, bottom) was measured by applying a 2-s P1 pulse to –50 mV from a holding potential (HP) of –100 mV. After a return to the HP for progressively increasing time intervals, a 1-s P2 pulse to +50 mV was applied. The time course of the progressively increasing peak P2 currents at +50 mV was then used to quantify the kinetics of recovery from CSI developed during the 2-s P1 pulse to –50 mV. Analysis of the 1-s i relationship is described in greater detail in the RESULTS, and additional methodological and analytical details are given in the appropriate sections of the text or figures.
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0.01) was determined using ANOVA. In figures, all data points are mean values ± SE obtained from the indicated numbers (n) of oocytes. |
RESULTS |
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Activation. For WT, R290A, R293A, and R296A, the a4 relationship could be estimated directly using a standard saturating tail-current protocol (22). Consistent with positive charge removal, all three mutants increased estimated k values (Fig. 2). Compared with WT, R290A produced a significant hyperpolarizing shift in a4 (approximately –30 mV), R293A was not shifted significantly, and R296A resulted in a significant depolarizing shift (approximately +30 mV). R290A thus stabilized channel open state(s), whereas R296A destabilized this state(s).
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R296A < R302A, and 2) estimates of activation time constants (
act,+50 mV) followed the same order (Fig. 3A). Similar to previous analyses of WT Kv4.3 channels (22, 30), estimates of act,+50 mV for R290A, R293A, and R296A could be fit with a sigmoidal a4 formulation (Fig. 3A, inset). Because of the overall smaller peak current amplitudes for R302A, act,+50 mV values were estimated from single-exponential fits to the approximate last one-quarter of the rising phase of the currents (thus the degree of sigmoidicity was not determined).
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A mutant slowed Kv4.3 activation kinetics, with the degree of slowing increasing as individual mutations "moved down" from the NH2 (extracellular) to COOH (intracellular) terminal of S4. Deactivation. Deactivation kinetics (single-exponential fits) were measured at –100 mV (details provided in Fig. 3), a potential where the value of a4 was zero for all expression conditions. This eliminated possible interpretative complications due to shifts in the steady-state activation relationship. Although each mutant altered deactivation kinetics rather than paralleling the effects of activation, two general, but opposing, effects were noted: R290A, R293A, and R296A all significantly slowed deactivation, whereas R302A significantly accelerated it (Fig. 3B).
Modifications in permeable cation species and concentrations have been reported to alter Kv4 channel gating kinetics (16, 21). To examine this possibility in detail, we investigated the mean reversal potential (Erev) of each RA mutant compared with the WT. We found that the three NH2-terminal mutants (R290A: –68.7 ± 1.7 mV, n = 16; R293A: –72.1 ± 2.5 mV, n = 9; and R296A: –68.0 ± 1.6 mV, n = 7) did not differ significantly from WT (–69.4 ± 0.9 mV, n = 9). By applying constant field assumptions (with intracellular K+ concentration = 100 mM and intracellular Na+ concentration = 15 mM), mean Erev values corresponded to estimated Na+/K+ permeability ratios (PNa/PK) of 0.039–0.050. The effects of these three mutants thus were not dependent on alterations in Kv4.3 selectivity or permeation properties. COOH-terminal R302A was found to depolarize Erev (–45.1 ± 3.1 mV, n = 11), corresponding to an estimated PNa/PK of 0.158. At present, we cannot exclude the possibility that changes in R302A selectivity and/or permeation properties may be a contributing factor to the kinetic changes noted in this mutant.
Macroscopic inactivation. We next determined the voltage dependence of the i relationship. From an initial HP of –100 mV, both WT and R302A peak current amplitudes elicited during a 1-s P2 pulse to +50 mV were not progressively reduced by a series of 1-s P1 pulses (5-mV steps from –95 to –80 mV), nor were peak P2 current amplitudes (+50 mV) increased by applying a 1-s hyperpolarizing pulse to –120 mV (WT: n = 4 and R302A: n = 3; data not shown). These results indicated that for WT and R302A, the steady-state value of i was 1.0 at a HP = –100 mV (refer to Fig. 5).
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A mutants promoted significant CSI at –100 mV, whereas COOH-terminal R302A did not alter this process.
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A mutants, a 1-s hyperpolarizing prepulse to –150 mV was applied to allow for (at least partial) recovery from inactivated closed states existing at –100 mV. This prepulse was then immediately followed by a series of 1-s depolarizing P1 pulses from –145 to +50 mV. Thus, the i relationships obtained for the first three R A mutants were isochronally dependent on 1-s prepulses to –150 mV and should not necessarily be viewed as steady-state relationships. For WT and R302A, since the steady-state value of i was 1.0, 1-s depolarizing voltage-clamp pulses ranging from –95 to +50 mV were applied directly from a HP of –100 mV.
Utilizing these protocols, mean 1-s i relationships could be obtained for all expression conditions (Fig. 5). For the purposes of the present analysis, all i curves were fit with single Boltzmann relationships. Compared with WT, each mutant produced shifts in mean V1/2 values, with the effects falling into two general, but opposing, categories. R290A, R293A, and R296A produced significant hyperpolarizing shifts in i (R290A > R293A R296A), whereas R302A produced a large depolarizing shift. The S4 NH2-terminal mutants thus promoted inactivated closed states, whereas COOH-terminal R302A stabilized noninactivated closed states.
In contrast to their effects on i, the RA mutants produced smaller and more subtle effects on the kinetics of macroscopic inactivation at +50 mV (a potential where the mean value of i was 0.0 for all expression conditions, thus eliminating interpretative complications due to shifts in the steady-state inactivation relationship). The effects again fell into two general, but opposing, categories: R290A, R293A, and R296A all produced an overall net slowing in the rate of macroscopic inactivation, whereas R302A produced no significant changes (Fig. 6A). For all expression conditions, inactivation at +50 mV could be well fit as a double-exponential process (Fig. 6B; see also Ref. 22). Overall, the fast and slow time constants of inactivation were either not significantly altered or minimally slowed; however, the relative value of the initial amplitude of the fast component of inactivation was significantly reduced in R290A, R293A, and R296A (Fig. 6C). Thus, the overall slowing in macroscopic inactivation kinetics produced by these three mutants was largely due to a reduction in the relative contribution of the fast component of inactivation, an effect not produced by R302A.
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10x for R290A and R293A and 100x for R296A), whereas R302A significantly accelerated it (10x; Fig. 7). This last mutant also produced an "overshoot" phenomenon in the amplitude of the recovering current compared with the P1 peak (Fig. 7, A and B). While the basis for this overshoot is presently unclear, no such phenomena were observed for the other current phenotypes. These results demonstrated that charge removal at R296 and R302 altered recovery kinetics over a time scale of approximately three orders of magnitude, a striking effect for the residues being in such tight spatial proximity.
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CSI.
Due to the markedly different effects of R296A and R302A on both i and kinetics of macroscopic recovery, we next specifically analyzed the effects of these two mutants on the development of and recovery from CSI compared with WT. For all three expression conditions, at –50 mV, the value of a4 was either equal to zero (WT and R296A) or no measurable net outward current could be observed (R302A; Fig. 2). There were, however, significant differences in the mean isochronal values of i (Fig. 5). If these values were due to alterations in the relative degree of CSI, then differences in both the extent and kinetics of development of CSI at –50 mV should have been observable. Consistent with this prediction, both WT and R296A developed significant CSI at –50 mV (WT: 95.8 ± 1.1% relative inactivation, n = 5; and R296A: 95.7 ± 1.1%, n = 6), whereas minimal (8.2 ± 5.5%, n = 2) to no (n = 4) CSI was observed for R302A (Fig. 8A). The kinetics of development of CSI were also significantly faster for R296A [time constant of CSI development (CSI,–50 mV) = 187.6 ± 16 ms, n = 8] than for WT (CSI,–50 mV = 544 ± 40 ms; n = 5; Fig. 8C, top).
Due to methodological limitations, it was not possible to measure recovery kinetics from CSI at a fixed potential where i was equal to 1.0 for all three expression conditions (see above). Therefore, relative measurements at a HP of –100 mV were made (Fig. 8, B and C, bottom). For WT, recovery from CSI (developed during a 2-s pulse to –50 mV) was sigmoidal [most likely indicating multiple inactivated closed-states (16)] and relatively fast [time constant of CSI recovery (CSI,rec,–100 mV) = 171 ± 8 ms, n = 5]. In contrast, R296A displayed apparently nonsigmoidal and significantly slower CSI recovery kinetics (CSI,rec,–100 mV = 4, 233 ± 498 ms, n = 7; Fig. 8, B and C, bottom). When present, recovery from CSI for R302A was quite rapid (CSI,rec,–100 mV = 58.4 ± 18.4 ms, n = 2; data not shown).
Thus, while the slower kinetics of recovery from CSI of R296A measured at –100 mV could not be separated unambiguously from effects due to shifts in i, they were nonetheless consistent with overall stabilization of inactivated closed states. Since i was equal to 1.0 at –100 mV for both WT and R302A, the effects of R302A could be attributed to genuine alterations in inherent Kv4.3 CSI kinetics resulting from destabilization of inactivated closed states.
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DISCUSSION |
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Role of S4 arginine residues in Kv4.3 CSI and recovery from CSI.
The molecular and biophysical mechanisms governing CSI and open-state inactivation have yet to be definitively characterized in any Kv4 channel (6, 16, 21). Our results therefore provide important insights for the understanding of Kv4 voltage-dependent gating transitions. The effects of the Kv4.3 S4 RA mutants support the proposal that, in the WT channel, the three NH2-terminal arginine residues (R290, R293, and R296) stabilize noninactivated closed states at resting membrane potentials. In contrast, COOH-terminal R302 stabilizes inactivated closed states at these potentials. Alterations of interactions involving these residues may have a significant impact on IA/Ito,fast function in the nervous and cardiovascular systems.
At present, the most widely accepted Kv4 channel gating models are based on two key underlying assumptions: 1) that the inactivated open state is nonabsorbing, and 2) that the voltage dependence of the final closed to open state transition is reverse biased (3, 6). These models propose that during maintained depolarization, channel open states (both conducting and transiently inactivated) will return to closed states and subsequently undergo absorbing inactivation. CSI would thus be the predominant mechanism responsible for inactivation at all potentials. Based on this model, it would be predicted that any maneuver to promote or stabilize inactivated closed states should result in an overall acceleration of macroscopic inactivation kinetics at depolarized potentials. However, our results indicate that stabilization of Kv4.3 inactivated closed states by R290A, R293A, and R296A produced only minimal effects on macroscopic inactivation kinetics at +50 mV [the net slowing was primarily due to a reduction in the relative amplitude of the fast component of inactivation (Fig. 6)]. They are thus difficult to reconcile with assumptions based upon these prior models (3, 6). Our results do suggest, however, that at depolarized potentials Kv4.3 inactivation possesses a significant open state component that is absorbing (22, 29).
Using kinetic analysis and Markov modeling, Wang et al. (29) proposed that absorbing open state inactivation is a dominant component in Kv4.3 channel gating. These investigators suggested an 11-state Markov gating model in which inactivation possesses multiple closed- and open-state pathways, with absorbing inactivation proceeding primarily from the open state. In contrast to previous Kv4 models (see above), this model specifically suggests that a direct transition between closed and open-inactivated states exists, a suggestion our laboratory proposed previously (22). While limitations with respect to recovery kinetics exist (discussed below), the model (29) accurately reproduces overall Kv4.3 CSI and open-state inactivation characteristics.
All S4 NH2-terminal mutants significantly slowed recovery and deactivation, whereas COOH-terminal R302A accelerated the two processes (although the latter effect was complicated; Fig. 7). These paralleling effects are consistent with the proposal that recovery from inactivation is coupled to deactivation (21, 22, 29, 30). Such coupling could account for the apparent discrepancy between Kv4.3 voltage-independent inactivation kinetics at depolarized potentials (where the rate constants for activation and inactivation display saturation) and voltage-dependent recovery kinetics at hyperpolarized potentials (where the rate constants for deactivation display voltage dependence) (21, 22, 30). Wang et al. (29) demonstrated that previous Kv4 gating models (3, 6) failed to adequately reproduce the experimentally measured voltage dependence of macroscopic recovery kinetics. While these investigators admitted that their gating model also fails in this respect, they have suggested that this limitation results from failure of their model, as well as those prior, to adequately account for the energetics of recovery-deactivation coupling. Our results strongly suggest that Kv4.3 recovery is indeed coupled to deactivation. We also support the proposition by Wang et al. (29) that the multiple components of Kv4.3 inactivation display different voltage dependencies. This is most notably evidenced by the apparent uncoupling of CSI from activation produced by R296A. As a result of the above findings, it is clear that new and verifiable Kv4 gating models are required, with our results providing a logical basis for the development of such models.
We propose that R290A, R293A, and R296A all slow recovery by slowing deactivation and stabilizing inactivated closed states. Conversely, R302A accelerates recovery by accelerating deactivation and stabilizing noninactivated closed states. The net effects of the mutations on recovery would thus arise from residue-specific variations in the proportion of inactivated closed states and alterations in deactivation kinetics. These combined effects appear to be particularly dependent on a stretch of only six amino acids (R296 to R302), wherein individual arginine charge elimination alters the time constants of recovery over a range of approximately three orders of magnitude.
One possible interpretative limitation to this conclusion should be noted. The acceleration in recovery kinetics for R302A could, theoretically, be separated from effects produced by shifts in a4. Due to restrictions on our recording protocols, however, such separation was not possible for R296A. Therefore, the inherent slowing of recovery kinetics noted for this mutant may not be as large as those recorded at a HP of –100 mV. Despite this limitation, the hyperpolarized shifts in i combined with slowing of recovery (both from macroscopic inactivation and CSI) are consistent with the three NH2-terminal mutants having induced significant stabilization of inactivated closed states at –100 mV.
Both we and others have previously hypothesized that Kv4.3 inactivation (both open and closed state) is sequentially coupled to activation (see Refs. 2 and 12). While the combined effects on a4 and i relationships of R290A, R293A, and R302A were not inconsistent with general predictions of sequential coupling, R296A produced unique effects in that a4 was significantly depolarized (Fig. 2), whereas i was simultaneously hyperpolarized (Fig. 5). R296A thus apparently produced sequential uncoupling of CSI from activation. It is important to note that in Shaker channels, the apparent voltage dependence of rapid open-state inactivation arises from coupling to activation (1, 31). Thus, the effects of R296A suggest that Kv4.3 CSI possesses intrinsic voltage dependence. In the absence of any evidence to the contrary, we still suggest that Kv4.3 open-state inactivation remains obligatorily coupled to activation (21, 22, 30).
Potential structural basis of S4 mutant effects. What could be the structural basis for our observations? In the absence of high-resolution structural data for any Kv4 channel, we used the crystal structure of the open-state conformation of Kv1.2 (18). Mapping of the Kv4.3 S4 sequence onto the corresponding structure of S4 for Kv1.2 (Fig. 9) indicated that, in the open state, R290 would be facing either a lipid or proteinaceous environment provided by the membrane bilayer or S5 of the adjacent subunit, respectively; R293 and R296 would be facing a proteinaceous environment provided by S1, S2, and/or S3; and R302 would be located close to the intracellular S4–S5 linker, potentially facing a proteinaceous environment provided by S2 or S3. The slowing of Kv4.3 recovery produced by the S4 NH2-terminal mutants could thus result from stabilization of hydrophobic interactions (R290A) or disruption of electrostatic and/or steric interactions (R293A and R296A) with residues in NH2-terminal S2 or along the length of S3. The acceleration of recovery produced by COOH-terminal R302A may correspond to removal of a rate-limiting step in recovery from CSI, possibly due to electrostatic and/or steric interactions of the residues with S2/S3 or other intracellular domains unresolved in the Kv1.2 model. Further refinement of these hypotheses should be possible in future studies with the use of charge reversal and/or glutamine mutations, resolving the importance of alterations in charge versus size on the kinetic properties we have presented here.
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A mutant altered basic activation characteristics. Such alterations are consistent with removal of positive charge from the voltage sensor, and this is the first study to characterize these effects explicitly in any Kv4 channel. We then provided strong evidence that recovery from inactivation is coupled to deactivation, as both processes followed similar trends. We went on to show that the precise location of arginine residues within S4 played an important role in the degree of development of CSI and recovery from CSI. At resting membrane potentials, the three NH2-terminal residues stabilized noninactivated closed states in the WT channel. Conversely, COOH-terminal R302 was found to stabilize inactivated closed states. It is the interactions mediated by these residues that determine the availability of the channel to enter the open state upon membrane depolarization and to regulate the kinetics of recovery from subsequent inactivation. We also demonstrated that the development of CSI can be sequentially uncoupled from activation by R296A, specifically. Taken together, these results extend our understanding of Kv4.3 gating transitions and reiterate the functional independence of Kv4 channels compared with the Kv1 family. |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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) are shown. Mean a4 data points were fit with the following single fourth-order Boltzmann relationship: a4 = 1/{1 + exp[(V – V1/2)/k]}4, where V is the test potential (in mV), V1/2 is the potential (in mV) where inactivation (i) = 0.5, and k is the slope factor (in mV). Fit parameters were as follows: WT, V1/2 = –36 mV and k = 14.5 mV (n = 8); R290A, V1/2 = –69 mV and k = 19.1 mV (n = 6); R293A, V1/2 = –36 mV and k = 17.8 mV (n = 7); and R296A, V1/2 = –7 mV and k = 14.7 mV (n = 7).
