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Am J Physiol Cell Physiol 294: C1476-C1484, 2008. First published April 2, 2008; doi:10.1152/ajpcell.00479.2007
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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS

A Kv channel with an altered activation gate sequence displays both "fast" and "slow" activation kinetics

Alain J. Labro,1 Alessandro Grottesi,2 Mark S. P. Sansom,2 Adam L. Raes,1 and Dirk J. Snyders1

1Laboratory for Molecular Biophysics, Physiology, and Pharmacology, Department of Biomedical Sciences, University of Antwerp, Belgium; and 2Laboratory for Molecular Biophysics, Department of Biochemistry, University of Oxford, Oxford, United Kingdom

Submitted 11 October 2007 ; accepted in final form 26 March 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The Kv1–4 families of K+ channels contain a tandem proline motif (PXP) in the S6 helix that is crucial for channel gating. In human Kv1.5, replacing the first proline by an alanine resulted in a nonfunctional channel. This mutant was rescued by introducing another proline at a nearby position, changing the sequence into AVPP. This resulted in a channel that activated quickly (ms range) upon the first depolarization. However, thereafter, the channel became trapped in another gating mode that was characterized by slow activation kinetics (s range) with a shallow voltage dependence. The switch in gating mode was observed even with very short depolarization steps, but recovery to the initial "fast" mode was extremely slow. Computational modeling suggested that switching occurred during channel deactivation. To test the effect of the altered PXP sequence on the mobility of the S6 helix, we used molecular dynamics simulations of the isolated S6 domain of wild type (WT) and mutants starting from either a closed or open conformation. The WT S6 helix displayed movements around the PXP region with simulations starting from either state. However, the S6 with a AVPP sequence displayed flexibility only when started from the closed conformation and was rigid when started from the open state. These results indicate that the region around the PXP motif may serve as a "hinge" and that changing the sequence to AVPP results in channels that deactivate to a state with an alternate configuration that renders them "reluctant" to open subsequently.

voltage-gated potassium channel

VOLTAGE-GATED K+ CHANNELS (Kv) respond to changes in the transmembrane voltage by opening or closing their ion-conducting pore. A functional K+ channel consists of a tetramer of {alpha}-subunits that surround a central ion-conducting pore. Each {alpha}-subunit has six membrane-spanning helices (S1–S6) with a pore loop (P) between S5 and S6. The detailed crystal structure of a voltage-independent bacterial K+ channel, KcsA, has resolved the molecular mechanism of K+ selectivity and permeation (14, 36, 51). The crystal structure of the mammalian voltage-gated Kv1.2 channel demonstrated that the structure of the ion-conducting pore (S5-pore loop-S6) is similar to that of KcsA (31, 32). Thus the molecular mechanism of K+ permeation is conserved between KcsA and Kv channels.

The opening and closing of the ion-conducting pore (gating process) of voltage-gated channels is triggered by a change in the membrane potential. The cytoplasmic activation gate of Shaker-type channels [of which hKv1.5 is a human homolog (44)], which open or close the ion-conducting pore, is located in the inner section of the ion permeation pathway. It has been proposed that residues V478 and F481 in Shaker (V516 and F519 in hKv1.5, respectively) occlude the pore in the closed state (11, 16). However, it remains controversial which residues form the gating "hinge" or pivoting point. Extrapolation of the predicted gating mechanisms of the Kv channel from archeabacterium Aeropyrum pernix (KvAP), KcsA, and the Ca2+-gated K+ channel from Methanobacterium autotrophicum (MthK) to Shaker suggested that G466 in Shaker (G504 in hKv1.5; Fig. 1) would act as hinge or pivoting point (24–26, 49). However, the crystal structure of bacterial K+ inward-rectifying channel (KirBac1.1) (28) suggested that G143 (the equivalent of P475 in Shaker and P513 in hKv1.5) was the pivoting point for gating, whereas the former more central-located glycine in the transmembrane helix G134 (corresponding to G466 in Shaker) was more important for packing. In human ether-a-go-go-related gene K+ channels (hERG), both the central and lower-located glycine in S6 are also required for tight protein packing, since the S6 helix is already inherently flexible (17). A substitution scan in K+ inward-rectifying (Kir1.1) channels showed that both glycine residues facilitate gating in a similar manner but that they are not absolutely required for pH-dependent gating of this class of channels (40).


Figure 1
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  		Fig. 1. Sequence of the lower S6 part of channels with an altered tandem proline motif (PXP). The residue numbering of human (h) voltage-gated K+ channel 1.5 (Kv1.5) is represented on top and the Shaker numbering at bottom in parentheses. Right: structure of the pore segment of a Kv1.2 subunit (31) with the S6 transmembrane helix in black. The structure shows that the S6 helix is bent around the well-conserved proline motif that is in the sequence alignment indicated in bold. The first proline is absolutely conserved in all Kv channels and the second only in channels that form functional homotetramers. The mutants created in this study were named according to their altered sequence, i.e., the double mutation P511A + V514P resulted in a AVPP sequence (with residue 511 indicated in bold); the latter name is used throughout this paper.

 
Substitution scan studies of the centrally located glycine residue in Shaker (G466) support that this glycine is important for channel biogenesis and may act as gating hinge (13, 34). A recent study in Kir3.4 channels further evidenced of the role of this central-located glycine in channel gating but underscored even more the role in protein packing by identifying that a small side chain is required at this position to prevent the interaction with other residues from, i.e., the selectivity filter (39). This study showed that, in Kir3.4 channels, "hinging" occurs at the residue immediately preceding this centrally located glycine (39). On the other hand, KvAP, KcsA, MthK, and KirBac1.1 lack a tandem proline motif (PXP) in the lower part of S6 that is conserved within the Kv1–4 families of voltage-gated (Shaker-type) K+ channels. Our results and those of others indicated that this motif may disrupt or bend the {alpha}-helical configuration of S6 (10, 29, 48). The crystal structure of the voltage-dependent rat (r) Kv1.2 channel showed that hinging could occur in the vicinity of the central glycine, but the structure also showed that the S6 helix is indeed bent around the PXP motif (31) (Fig. 1). Apparently this {alpha}-helical disruption is crucial for voltage-dependent gating of this class of channels, but the proline-containing motif does not need to be located at the evolutionary conserved position (29, 48). Indeed, hKv1.5 channels with an altered PXP sequence, PVAV, AVPP, or AVAP (Fig. 1), were functional but displayed marked changes in channel gating. These channels activated very slowly, and the voltage dependence of activation did not reach saturation during 5-s voltage steps up to +110 mV (29). Here we report that channels with a AVPP sequence display two gating modes.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Molecular Biology

Starting from hKv1.5 cDNA cloned in a pBK-CMV vector, mutations were introduced in hKv1.5 with the QuikChange Site-Directed Mutagenesis kit (Stratagene, La Jolla, CA) using mutant primers. After the PCR-based mutagenesis, a Pst1-BamH1 fragment containing the mutation was cut out of the PCR-amplified vector and ligated in hKv1.5/pBK-CMV to replace the wild-type (WT) sequence. Double-stranded sequencing of the exchanged fragment and the adjacent sequence confirmed the presence of the desired modification and the absence of unwanted mutations.

Cell Culture and Transfection

Plasmid DNA for mammalian expression was obtained by amplification in XL2 blue script cells (Stratagene). The plasmid DNA was isolated from the bacterial cells with the GenElute HP plasmid maxiprep kit (Sigma, St. Louis, MO). The cDNA concentration was determined with ultraviolet absorption. Mouse Ltk– cells were cultured in DMEM medium supplemented with 10% horse serum and 1% penicillin/streptomycin. The culture was passed every 3–4 days. For experiments, the cells were transfected with 0.5–5 µg of WT or mutant subunit cDNA. Transfections were done according to the lipofection method using lipofectamine (Invitrogen, Carlsbad, CA). After transfection (12–24 h), the cells were trypsinized and used for analysis within 12 h. A stable cell line was created for the mutant with an AVPP sequence as described previously (43). For this purpose, the mutant AVPP was cloned in a pMSV-neo vector with an inducible promoter. For experiments, channel expression was induced by adding 100 µl dexamethasone (of a 100 µM stock solution) to the cell medium 16–24 h before patch-clamp analysis.

Electrophysiology

Current recordings were made with an Axopatch-200B amplifier (Axon instruments, Foster City, CA) in the whole cell configuration of the patch-clamp technique. Experiments were done at room temperature (20–23°C); current recordings were low pass filtered and sampled at 2–10 kHz with a digidata 1200A data acquisition system (Axon instruments). Command voltages and data storage were controlled with pClamp8 software (Axon Instruments). Patch pipettes were pulled from 1.2 mm kwik-fill borosilicate glass capillaries (World Precision Instruments, Sarasota, FL) with a P-2000 puller (Sutter Instruments, Novato, CA). After heat polishing, the resistance of the patch pipettes was <3 M{Omega}. The cells were perfused continuously with a bath solution containing (in mM) 130 NaCl, 4 KCl, 1.8 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, adjusted to pH 7.35 with NaOH. The pipettes were filled with intracellular solution containing (in mM) 110 KCl, 5 K4BAPTA, 5 K2ATP, 1 MgCl2, and 10 HEPES and was adjusted to pH 7.2 using KOH. Junction potentials were zeroed with the filled pipette electrode in the bath solution. Experiments were excluded from analysis if the voltage errors originating from series resistance exceeded 5 mV.

The holding potential was –80 mV unless specified otherwise. The voltage protocols were adapted to adequately determine the different biophysical parameters of mutant channels. For WT hKv1.5, a 900-ms voltage ramp from –80 to +100 mV was optimal. For AVPP, a 1,200-ms ramp from –80 to +100 mV was used, since ramps longer than 1,200 ms were blurred by the "slow" activation state. Because of the Goldman-Hodgkin-Katz (GHK)-type rectification of hKv1.5 (43), the current recordings from ramp protocols were normalized with the GHK current equation;

Formula 1(1)
where P(o) represents the normalized open probability, Ik the current measured, V the applied voltage, Ek the reversal potential of –80 mV with the solutions used, [K]i the intracellular K+ concentration (4 mM in our case), and R, T, and F have their usual meaning (8). Time constants of activation and deactivation were determined by fitting the current recordings with a single exponential function. The voltage dependencies of channel activation were fitted with a Boltzmann equation: y = 1/{1 + exp[–(V – V1/2)/k]}, in which k represents the slope factor, V the applied voltage, and V1/2 the voltage at which 50% of the channels are activated. Results are expressed as means ± SE, and n the number of cells analyzed.

Molecular Dynamics Simulations

A series of simulations (each 10 ns long) of the isolated S6 helix of hKv1.5 K+ channel, modeled on the closed and open conformation, was carried out in the presence of a mixed 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine/1-palmitoyl-2-oleoyl-sn-glycero-3-phosphorylglycerol bilayer. The following systems have been set up: WT hKv1.5 and the PXP motif variants AVAP and AVPP. The initial protein coordinates for the closed conformation were modeled using Modeller and PDB entry 1K4C [PDB] as a template. The coordinates for the open conformation were extracted from the rKV1.2 X-ray structure (PDB entry: 1P7B [PDB] , www.rcsb.org). The transmembrane S6 helix was defined as extending from residue Val491 to Thr527, the PXP motif encompassing residue 511–513. The system (i.e., protein plus membrane) was solvated with SPC water molecules (4), retaining all the crystallographic waters.

Molecular dynamics (MD) simulations were performed with GROMACS 3.1.4 (30) (www.gromacs.org) with a modified version of the GROMOS-87 force field (47). Lipid parameters were based on those by Berger et al. (5) and Marrink et al. (35). For the lipid-protein interactions, the GROMOS parameters were used. Simulations were carried out at constant pressure and temperature, NPT statistical ensemble, with periodic boundary conditions. The initial velocities were taken randomly from a Maxwellian distribution at 300 K. The temperature was held constant by coupling to an external bath (22). Long-range electrostatic interactions were calculated using the Particle Mesh Ewald summation methods (9). Lennard-Jones interactions were calculated using a cutoff of 0.9 nm. The pair lists were updated every 10 steps. The LINCS algorithm (20) was used to constrain bond lengths. The time step was 2 fs, and coordinates were saved every 0.1 ps. Principal components analysis was performed as previously described to (1, 15). The computational Markov modeling is described in the supplemental data (Supplemental data for this article is available online at the American Journal of Physiology: Cell Physiology website.).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
We have shown previously that at least one proline is required in the lower part of S6 for a functional hKv1.5 channel (29). When the first proline of the evolutionary conserved motif was replaced by an alanine P511A (i.e., a AVPV sequence), no time-dependent currents were detected within a voltage range of –130 to +130 mV despite the presence of the channel in the plasma membrane. Note that the hKv1.5 numbering used here is the revised one (accession no. P22460) and that the P511A channel mutant in fact corresponds to the previously described P509A mutant (29) that was named according to the original numbering by Tamkun et al. (44). As expected, promoting a complete {alpha}-helix by substituting alanine residues for both prolines (P511A + P513A) also resulted in a nonfunctional channel. However, both mutants could be rescued by the introduction of a proline at a position different from the conserved one. The double mutant P511A + V514P resulted in an AVPP sequence (Fig. 1) and rescued the P511A mutation, since voltage-dependent K+ currents were observed after patch disruption. Upon the first depolarization to +60 mV from a holding potential of –80 mV, the current activated quickly with time constants in the millisecond range and subsequently inactivated slowly (Fig. 2C). This time course of current activation was similar to WT hKv1.5 (Fig. 2A) and to that of the V514P mutation in WT, resulting in a PVPP sequence (Figs. 1 and 2B). However, during the second depolarization to +60 mV, the current activation was markedly slowed (s range) and displayed no time-dependent inactivation (Fig. 2C). This second type of activation (Fig. 2D) was similar to that of channels with an AVAP (triple mutant P511A + P513A + V514P; Fig. 2E) and a PVAV sequence (mutant P513A, Fig. 2F), all displaying markedly altered channel gating.


Figure 2
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  		Fig. 2. Currents of hKv1.5 channels with an altered PVPV motif. The voltage protocol is shown above the resulting current tracings; notice the differences in scale bars in A–F. Horizontal bar on left indicates the zero current level. A: wild-type (WT) hKv1.5 with a PVPV sequence displaying fast activation followed by slow inactivation. B: control mutant V514P with a PVPP sequence displayed activating currents that resembled WT hKv1.5. C: mutant with an AVPP sequence: during the first depolarization the current activated quickly, followed by a slow decline indicative of slow inactivation (tracing indicated by "1"). During the second depolarization, the fast current component was absent; instead, a very slow-activating current component was observed (tracing indicated by "2"). D: family of currents for AVPP in its "slow" gating mode. E and F: currents of mutant channels with an AVAP and a PVAV sequence, respectively. The currents of these channels were comparable to those of AVPP in the slow mode (D).

 
The second gating mode that was observed in channels with the AVPP sequence was characterized by an apparent threshold of activation around –30 mV, comparable to WT hKv1.5 (PVPV). However, 5-s voltage steps up to +110 mV were insufficient to reach saturation of the activation (Fig. 3A). It is unlikely that this behavior was due to the proline mutation at position 514 itself, since this mutant V514P in WT hKv1.5 (PVPP) behaved similar to WT (PVPV), both in its voltage dependence and kinetics of activation (Fig. 3A). The activation time constants of the second gating mode of AVPP were ~200 times slower than WT (PVPV) or the control mutant V514P (Fig. 3B). The time constant of activation of AVPP in the initial gating mode was 10.9 ± 1.0 ms (n = 8) at +60 mV, which is approximately three times slower than WT or the control mutant V514P (PVPP) but markedly (~55 times) faster than that of its second gating mode (Fig. 3B). Throughout we will use "fast" and "slow" mode as a shorthand for the initial gating mode that is characterized by fast WT-like activation or the second gating mode that shows slow current activation, respectively. The subsequent channel inactivation of the fast mode had kinetics that displayed no or a limited voltage dependency and were similar to those of WT hKv1.5 (Supplemental Fig. S1). Approximating the fast mode inactivation of AVPP with a single exponential function yielded a time constant of 702 ± 46 ms (n = 8) at +30 mV. The deactivation time constant of AVPP in the slow mode was comparable to that of the control mutant V514P with a PVPP sequence (Fig. 3B); both were approximately six times slower than WT (PVPV). All mutants had a reversal potential around –80 mV (data not shown), indicating that the K+ selectivity of the channels was not altered.


Figure 3
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  		Fig. 3. Biophysical properties of WT (PVPV), PVPP, and AVPP. A: voltage dependence of activation of WT (PVPV) (circles), the control mutant PVPP (open squares), and AVPP slow mode (triangles). For WT and PVPP, the voltage dependence of activation reached saturation. A fit with a Boltzmann function yielded a midpoint of –14 ± 1 mV and a slope factor of 5.8 ± 0.2 mV (n = 9) for WT (PVPV). The activation curve of PVPP corresponded well to WT with a midpoint of –22 ± 2 mV and a slope factor of 6.4 ± 0.4 mV (n = 7). When AVPP was in its slow mode, the voltage dependence of activation did not reach saturation. The threshold for activation was –30 mV (similar to WT), but the activation curve had a midpoint of at least +50 mV and a slope factor of 32 mV (n = 5). B: time constants of activation and deactivation are shown for WT, PVPP, AVPP slow (triangle) and AVPP initial fast (inversed triangle) mode. PVPP was somewhat slower than WT (PVPV). The activation of the "fast" mode of AVPP at +60 mV was slightly slower than that of PVPP and ~3 times slower than WT. In contrast, AVPP slow mode was ~200 times slower than WT. The difference in gating kinetics between AVPP fast (inversed triangle) and slow (triangles) mode at +60 mV is indicated with an arrow. Notice the logarithmic time scale.

 
These observations suggested that the AVPP channel initially exists in a mode from which normal (fast) activation is possible but that a single depolarization/repolarization cycle was sufficient to trap the channel in a second slow mode. To test whether the switch to the slow mode was caused by the activation, deactivation, or inactivation, the duration and the voltage of the initial depolarizing step were altered (Fig. 4). Even short depolarization steps that did not induce significant inactivation were sufficient to trap the channel in the slow mode, suggesting that the switch was dependent on the activation or the subsequent deactivation but not inactivation (Fig. 4, A–C). After longer depolarization, the initial fast and the second slow mode reached the same steady-state current level with identical tail currents (Fig. 4A). With very short depolarization steps, a fraction of channels remained in the fast mode after the first step, presumably because not all channels had been activated (Fig. 4, D and E). Consequently, a fraction of channels still activated in the fast mode during the following depolarization. If activation caused the switch from one gating mode to the other, it should be possible to calculate the expected current amplitude for the first, second, and third step based on the activation time constant of the initial fast mode. To test this, we created a stable cell line in which the current density of the fast mode was 112 ± 11 pA/pF (n = 20) (Fig. 4F). Because the time constant of activation of the fast state was 10.9 ms at +60 mV, 8-ms depolarizing steps should activate only 52% of the channels. Indeed, the current amplitude upon the first 8-ms depolarizing step was 48 ± 7 pA/pF (n = 11), which is ~43% of the current density in this cell line. The expected values for the subsequent second and third depolarizing steps were 25 and 12% of the total value of 112 ± 11 pA/pF, being 28 ± 3 and 14 ± 1 pA/pF respectively. The measured values for this second and third step also compared very well to the expected ones (Fig. 4F). Furthermore, the sum of the current amplitudes obtained during all three depolarizing steps accounted for 90% of the activatable channels and therefore approached the current density value of this cell line (93 ± 15 pA/pF, n = 11; Fig. 4F). These results indicate that the activation (or deactivation) and not the subsequent inactivation trapped the channel in the slow mode; this trapping was observed at any depolarizing voltage above the apparent threshold (–10 mV; supplemental Fig. S2).


Figure 4
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  		Fig. 4. Trapping AVPP in the slow mode depends on channel activation. The duration of the depolarizing step to +60 mV after patch disruption was varied from 6 s (A) to 8 ms (E). A: during the initial 6-s depolarization step, the AVPP channel activated fast followed by slow inactivation (current trace 1). During the second depolarization, the activation was slow (trace 2). At the end of these long pulses, both tracings reached the same "steady" state. The tail currents, elicited by stepping to –30 mV, were also similar. In B and C, 200- and 50-ms depolarizing steps were applied. In both cases, a fast current activation was observed during the first depolarization (trace 1). Upon the second depolarization, the current activation was slow (trace 2), and the channel was already trapped in its slow mode. In D and E, the depolarizing step was decreased to 20 and 8 ms, respectively. Even these very short steps trapped the channel in the slow mode. However, with these short depolarizing steps, not all channels were activated during the first depolarization (trace 1), and a smaller fast current component remained upon the following depolarizations (traces 2–4). F: to determine that the remaining fast current component upon the second depolarization originated from channels that were not activated during the 1st depolarization, the expected current amplitude for the 1st, 2nd, and 3rd depolarizing step was calculated for 8-ms-long depolarizations (see text). A stable cell line was used of which the current density of the fast mode was 112 ± 11 pA/pF (n = 20, open circle). As such, the measured current amplitudes upon the 1st, 2nd, and 3rd step (8-ms depolarizations as in E) were compared with the expected ones. After leak correction and subsequent normalization to the cell capacity, the measured current densities were plotted as a function of the step number (n = 11, open squares). These values compared well with the values that were calculated (circles), based on the activation time constant at +60 mV and the mean current density (fast mode) of the stable cell line.

 
Because a single depolarization trapped the AVPP channel in the slow gating mode, the voltage dependence of activation for the fast mode could not be determined with the usual step protocols. Therefore, we used a voltage ramp (from –80 to +100 mV) to obtain an approximation of the voltage dependence of the fast mode. To validate this approach, this procedure was also used to determine the activation curve of WT channels (PVPV) (Fig. 5B). Given the (GHK-type) outward rectification of hKv1.5 (43), we normalized the evoked current with the Goldman-Hodgkin-Katz dependence on driving force (8, 21). This resulted in an activation curve with a midpoint of –7.7 ± 1.2 mV and a slope factor of 6.2 ± 0.3 mV (n = 7), which compared reasonably well with the "steady-state" "isochronal" activation curve with a midpoint of –14.3 ± 1.0 mV and a slope factor of 5.8 ± 0.2 mV (n = 9; Fig. 5, A and B). For channels with the AVPP sequence, we obtained an activation curve with a midpoint of +16.5 ± 0.8 mV and a slope factor of 7.9 ± 0.4 mV (n = 7), which represents a +24-mV positive shift compared with WT (Fig. 5C). Accounting for this shift in voltage dependence of activation, the kinetics of the fast mode determined at +60 mV (10.9 ± 1.0 ms, n = 8) were comparable to those of WT channels and those of the corresponding control mutant V514P (PVPP sequence) at +40 mV (4.2 ± 0.3 ms for WT n = 9 and 7.1 ± 0.9 ms for V514P n = 7).


Figure 5
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  		Fig. 5. Voltage dependence of activation of WT hKv1.5 (PVPV) and AVPP fast mode. A: isochronal activation curve of WT on the right was determined from normalized tail current amplitudes using the step protocol shown on the left. The solid line represents a Boltzmann fit with a midpoint of –14 mV and a slope factor of 6.2 mV. B: activation curve of WT determined with a voltage ramp. Left: 900-ms voltage ramp from –80 to +100 mV with the corresponding current below. Plotting the normalized permeability P(o) (see MATERIALS AND METHODS) as a function of the applied voltage resulted in an activation curve that is represented on the right. Fitting the activation curve between –30 mV and +40 mV (boundaries indicated by vertical bars) with a Boltzmann function yielded parameters close to the values obtained in A, i.e., a midpoint of –10 mV with a slope factor of 5.9 mV. Note that both protocols A and B were recorded within the same cell. C: determination of the voltage dependence of activation of AVPP initial fast mode. Left, current recorded with a 1,200-ms voltage ramp that was applied after patch disruption. Right, activation curve of the fast mode of AVPP obtained as in B. Fitting the curve with a Boltzmann function between –15 and +45 mV yielded a midpoint of +16 mV and a slope factor of 6.7 mV (solid line).

 
The initial fast mode of AVPP was not caused by an incomplete intracellular perfusion, since holding the cell at –80 mV for 10 min before the application of the first depolarizing pulse still resulted in fast current activation (data not shown). To examine whether the fast mode could be recovered, the cells were clamped for varying durations at voltages where the channels normally close (–40 mV and below). Holding the cell sufficiently long at negative voltages (>10 min) partially recovered the channel to the fast mode (Fig. 6A). Moreover, the activation time course of the recovered fast mode was similar to that of the initial one upon patch disruption, 8.4 ± 0.5 ms compared with 10.9 ± 1.0 ms at +60 mV (n = 8), respectively. To determine the recovery time constant, the current amplitude of the fast component was expressed as a function of the recovery interval (Fig. 6B). Approximating this relation with a single exponential function yielded a very slow recovery time constant of 400 ± 30 s at –80 mV (n = 5; Fig. 6C). Interestingly, there was no statistical difference to this recovery time constant in the voltage range from –40 to –80 mV (Fig. 6C), although the fraction of recovered channels in the fast mode was reduced at the most positive holding potential of –40 mV (supplemental Fig. S3).


Figure 6
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  		Fig. 6. Time-dependent AVPP channel recovery to the fast mode. A: voltage protocol with the corresponding currents. The first current trace recording after patch disruption is represented with 1, and the channel was activated in the fast mode. Upon the second depolarization, 15 s later, the channel was in the slow mode (trace 2). Holding the cell for longer time periods at –80 mV partially recovered the channel in the fast mode. After 20 min, ~40% of the channels were recovered in the fast mode, trace 9. The activation time constant during the first step after recovery (8.4 ± 0.5 ms at +60 mV) was similar to that of the initial one upon patch disruption (10.9 ± 1.0 ms, n = 8). B: the fast current amplitude from A (indicated by arrow on top) plotted as a function of the recovery time or interepisode time. The data from current traces 1, 2, and 9 from A are indicated. Note the increase of the fast current component with longer recovery periods at –80 mV. C: time constants of channel recovery to the fast mode at various holding potentials, as indicated in each panel. Data points represent the averages of individual data (leak corrected), each obtained as in B and normalized to the final fast current amplitude (corresponding to the current amplitude of trace 9 in A). The line in each panel illustrates a monoexponential fit to the average data points. The average time constant of recovery ± SE with the no. of cells in parentheses is represented in the right corner of each panel. The time-dependent channel recovery to the fast mode did not display any significant voltage dependence, since similar and statistically not different recovery time constants were obtained with holding potentials between –40 and –80 mV.

 
To test the effect of removing and/or shifting proline residues away from the PXP motif on the flexibility of the S6 {alpha}-helix, MD simulations were performed on the S6 transmembrane domain of WT hKv1.5 and the mutant channels AVPP and AVAP in a membrane-mimicking environment. For both WT and mutant channels, we ran three independent simulations starting from either a closed or open channel conformation, each time using a new set of initial velocities obtained at 300 K (see MATERIALS AND METHODS for details). Analysis of the stability and reliability of all simulations (based on calculation of root mean square deviation) showed that the isolated S6 helix of all systems was quite mobile and subjected to short time scale rearrangements (~1, 1.5 ns) (data not shown). Secondary structure content analysis was performed using the Dictionary of Secondary Structures of Proteins software (27 and Fig. 7). The analysis revealed that, in the simulations starting from the closed conformation (left part in Fig. 7), the {alpha}-helical content for WT was lost in the middle of the helix around residue 511, which corresponds to the first proline of the PXP motif. This is an indication for the presence of a hinge point around this position. For the AVAP variant, the loss of {alpha}-helical structure was more pronounced, and the COOH-terminal half of the helix adopted either a p-helix or turn conformation, which represents a different dynamic behavior for the AVAP mutant compared with WT. On the other hand, the AVPP mutant displayed an average {alpha}-helical content per residue similar to WT and displayed the marked loss of a structure and the gain of a turn-type conformation around residues 510–514 comparable to WT.


Figure 7
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  		Fig. 7. Secondary structure probability of the S6 helix of WT, AVAP, and AVPP. Secondary structure content was calculated using the Dictionary of Secondary Structures of Proteins software and shown as the average percentage of time spent by a given residue in the corresponding secondary structure. Left: closed helix conformation simulations. Right: relative to the open conformation. Red, {alpha}-helix; green, p-helix; gray, g-helix; yellow, random coil; brown, β-bend; blue, β-turn.

 
Analysis of the simulations carried out starting from the open S6 helix conformations (Fig. 7, right) revealed that the WT helix was quite stable with again the loss of {alpha}-helical content at position 511 that was clearly evident. This loss in {alpha}-helical content was absent in both the AVAP and the AVPP mutant. Interestingly, both showed a quite similar secondary structure content in the open helix simulations, suggesting that the open state of the S6 helix of these mutants is more stable than WT on this time scale.

A further indication of this trend is shown in the analysis of fluctuations performed by measuring the mean square displacement from the average structure according to the first principal components of the covariance fluctuations matrix. Figure 8 clearly shows that, for WT, in both closed and open conformations, there was a clearly higher displacement located near residue 509 over the entire time scale of the simulation. Because these fluctuations were focused around the PXP motif, this region of the helix was less stable and likewise more flexible. Starting from a closed conformation, the AVPP variant displayed a higher flexibility around the residues 510–511, a behavior qualitatively comparable to the WT counterpart. However, in the open helix conformation, the flexibility pattern according to the principal components suggested that the intrinsic dynamical behavior of AVPP matched that of AVAP, indicating that the movements of the isolated S6 helix differ from that of WT.


Figure 8
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  		Fig. 8. Mean square displacement in the S6 helix of WT, AVAP, and AVPP. Mean square displacement from average position per C{alpha} carbon atom, calculated by filtering the fluctuations along selected principal components. A peak in the graphs represents a point along the sequence where the flexibility of the corresponding C{alpha} carbon is high. Left: simulations performed in the closed S6 helix conformation. Right: corresponding displacement for the open helix simulations.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The evolutionary conserved tandem proline motif PVPV is essential for a functional Shaker-type hKv1.5 channel (29). Replacing both prolines by an alanine (promoting a complete {alpha}-helix) resulted in a nonfunctional channel; even replacing the first proline by an alanine, creating the sequence AVPV, resulted in a nonfunctional channel. However, both channel mutants reached the plasma membrane, as assessed by confocal microscopy (29). This was in agreement with the results for the corresponding mutant in Shaker (P473A) that displayed clear gating currents but no ionic current (16). However, the introduction of an extra proline at position 514 (mutation V514P), creating a AVPP sequence, rescued the nonfunctionality of P511A. This AVPP sequence resulted in a channel that existed initially in a fast gating mode (Figs. 2C and 4), but upon the first activation or deactivation the channel became trapped in a slow one (Fig. 2, C and D, and 3). The gating kinetics of this slow mode were comparable to those of channels with a AVAP or a PVAV sequence (Fig. 2, E and F). However, these AVAP and PVAV channels did not display two gating modes, since the current activation upon the very first depolarization was the same as the second (data not shown). The biophysical properties of these channels and of AVPP in the slow mode were profoundly altered compared with WT hKv1.5 (PVPV). Because very short depolarizing steps were sufficient for AVPP channels to switch to their slow gating mode (Fig. 4, D–F), it was presumably not the inactivation that caused the switch.

The fast gating mode of AVPP channels could be recovered, but this recovery was extremely slow (time constants in the order of 10 min) and voltage-independent for holding potentials between –50 and –80 mV (Fig. 6). Because of this very slow recovery process, we could not characterize the fast mode of AVPP completely. With the approximation of using a voltage ramp (Fig. 5C), the estimated voltage dependence of the fast mode was much closer to that of WT compared with that of the slow mode, which itself did not saturate (Fig. 3A). Furthermore, the activation time constant of the fast mode corresponded well to that of WT channels (PVPV) and the control mutant V514P with a PVPP sequence, especially if corrected for midpoint shifts. Remarkably, the deactivation time constants of AVPP fast and slow mode were similar and resembled those of the control mutant V514P, and both were markedly slower than WT (Fig. 3B). Such slowing of channel deactivation was also observed with alanine or isoleucine substitutions at this position (38). These V514A and V514I mutations displayed an isolated "deactivation" failure that was caused by stabilizing the open state while destabilizing the closed state. Such a slowing of deactivation is apparently common for mutations at this position given the results for the V514P mutant (PVPP) and the AVPP channel.

Previous results have suggested that the proline residues of the tandem proline motif (PVPV) disrupt the S6 {alpha}-helix (10, 29, 48). The question remained whether this putative disruption of the {alpha}-helix is required solely for structural packing of the channel protein or whether it also serves a functional role by creating a gating hinge. If the PXP motif disrupts ("bends") the S6 {alpha}-helix, this region should have greater degrees of freedom and could therefore generate flexibility. This idea is supported by molecular modeling that indicated that the PXP motif distorted the {alpha}-helix, creating a hinge region (6, 7, 33, 45, 46). If the PXP motif is important in WT to reorient the lower part of S6 (channel gate) during gate opening or closing, the same most likely accounts for the mutants. The biophysical data indicated that the Kv channel mutant with a AVPP sequence initially exists with the lower S6 region in a conformation that allows "normal" activation, but activation or subsequent deactivation locks S6 in a different conformation that precludes normal activation. MD simulations of the S6 helix of WT hKv1.5 revealed significant movements around the PVP motif in both the open and closed conformation. However, for the AVAP sequence, the simulations suggested that the helix is rigid in both the open and closed conformation, since the principal component analysis identified no specific movements (Fig. 8). The absence of clear movements may explain the slow kinetics of this AVAP mutant. Interestingly, the MD simulations of AVPP showed that this mutant displayed two different behaviors (as in the functional analysis). When starting from the open conformation, the S6 helix of the AVPP mutant was as rigid as AVAP, which could be correlated with the slow gating kinetics. In contrast, when starting from a closed conformation, AVPP displayed movements similar to WT, which suggests the presence of a flexible hinge and may explain the initial observed WT-like kinetics. This indicates that the AVPP sequence can either introduce flexibility in the S6 helix or be rigid, dependent on starting from a closed or open channel conformation, respectively. Because this switch in behavior occurs during channel gating, this indicates that this region (and presumably also the PVPV region in WT) undergoes conformational movements during channel opening or closing and could therefore act as a hinge.

Olcese et al. (37) described a double mutation in Shaker that converted the inactivated state into a second open state. One of these mutations, I470C (I508C in Kv1.5), is in the vicinity of the PXP motif. This raises the question whether the double-gating behavior of the channel with an AVPP sequence can be explained by a modified inactivated state. In this case, the effect of the AVPP mutation would not (only) be at the level of the S6 "bundle crossing" but also at the selectivity filter. Under this hypothesis, the reduced flexibility of the S6 helix with simulations starting from the open conformation would indicate that the cytoplasmic gate becomes locked in the open configuration after the first activation. The observed second slow gating mode would then be due to an alteration in the selectivity filter that converts from a new nonconducting state to the former "inactivated" state but which is now conducting. However, the inactivation kinetics of the fast mode are largely voltage independent and clearly slower than the activation time constants of the slow mode (supplemental Fig. S1). Furthermore, because the channel still deactivates, this would require that the normal-conducting (open) state of the selectivity filter becomes now a nonconducting one since the cytoplasmic gate is locked open and cannot seal off the channel pore anymore. Although we cannot completely rule out this possibility, it relies on several additional assumptions. Therefore, we prefer the proposal that the slow mode is triggered when the cytoplasmic S6 gate closes into a configuration that precludes fast activation.

Computational models to simulate the AVPP behavior indicated that a switch in gating mode at the level of an intermediate closed state matched the experimental data best (Supplemental Fig. S4).


Formula 2

(2)

If the switching is located closer to the open state, the behavior during depolarization can still be simulated, but not the relatively fast deactivation combined with slow activation kinetics (Supplemental Fig. S4). The observation that the deactivation after the first (fast) and subsequent (slow) activation proceeds with the same time course can be interpreted that the initial closing step(s) is the same in both gating modes but that a further reorientation upon deactivation shifts the S6 gate in a conformation from which subsequent activation is slow. Thus, if we approach the behavior of the AVPP mutant with a simplified version of the 15 or 16 state model (50), the state transitions of one subunit of the tetramer would schematically be as follow.


Formula 3

(3)

In equations 2 and 3, the A to O transition refers to the final concerted step of opening and presumably involves a movement(s) of the complete tetramer. Computational modeling suggested (Eq. 2, Supplemental Fig. S4) that the channel switches to a reluctant closed state (C*) at an intermediate closed state (Ci). This C* branch can thus be seen as a mirror of the normal closed C branch. The recovery process from this reluctant closed state C* to the normal C state is then extremely slow. If we combine this equation 2 with the results of the MD simulations, the normal "WT" closed C state is defined by a S6 segment that shows flexibility around the PVP motif and allows both fast channel activation and deactivation from and to this state or configuration. The reluctant C* branch on the other hand is characterized by a S6 segment that is rigid and results in slow gate opening. The results of the MD simulations are in good agreement with this interpretation, since the S6 helix revealed flexibility starting from the closed conformation but not when the channel was in the open state. Thus once the open or preopen activated conformation is reached, the S6 segment becomes less flexible, and the channel switches to the reluctant C* branch upon channel closure.

The cytoplasmic activation gate of Shaker-type channels is formed by the lower part of S6 (11, 12, 16, 38). In channels with a AVPP sequence, this lower part of S6 switches between a nearly normal fast conformation and an altered slow conformation. Although we cannot exclude the possibility that another associated channel segment locked the cytoplasmic activation gate in this new position, it is likely that the altered gating hinge reoriented and locked the channel in a slow mode. Presumably, channels with a PVAV or AVAP sequence instead of WT (PVPV) exist in a similar but permanent slow conformation, since MD simulations showed lack of flexibility in both conditions. One could speculate that the association with the KCNE1 subunit results in a similar permanent slow conformation in the KCNQ1 subunits, resulting in the known slow activation kinetics of the cardiac slow component of the delayed-rectifier current (IKs) (2, 42). Similarly, N-type and P/Q-type calcium channels switch from a rapidly activating "willing" (W) mode to a more-difficult-to-activate reluctant (R) mode upon binding of Gβ{gamma} subunits (3, 18, 23, 41). These channels can be shifted back from the R mode toward the W mode by stronger depolarizations or by protein kinase C phosphorylation. Apparently this R mode and its different modulation is an intrinsic property of the calcium channel itself (19). This indicates that, similar to our hKv1.5 channel with an AVPP sequence, these calcium channels reversibly switch between a fast activating W mode and a slow or R mode.

In conclusion, our results indicate that changing the evolutionary conserved PVPV motif in hKv1.5 into AVPP resulted in a fully folded channel that resides in the membrane and reversibly switches between a fast and a slow gating mode. This indicates that the region around the double proline (PXP) sequence reorients upon channel activation and/or deactivation. MD simulations of the isolated S6 helix show that the AVPP sequence can introduce flexibility in this region (fast mode) but at the same time being very rigid (slow mode). Because only a minor alteration around the crucial PXP motif may lead to faster or slower channel gating, the large variety in gating kinetics observed throughout the Kv family of channels may result from only subtle differences in the gating machinery that transduces voltage sensing to channel opening.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by grants from the Fonds voor Wetenschappelijk Onderzoek Vlaanderen (FWO-G008404, FWO-G015206, and FWO-1504407N) and GOA-University of Antwerp.


   ACKNOWLEDGMENTS
 
We thank T. Bruyns and T. de Block for technical assistance in the molecular biology. A. J. Labro is a postdoctoral fellow with the "Fonds voor Wetenschappelijk Onderzoek Vlaanderen."

Current address for A. Grottesi: Consorzio Interuniversitario per le Applicazioni del Supercalcolo per Universita e Ricerca, Rome, Italy.


   FOOTNOTES
 

Address for reprint requests and other correspondence: D. Snyders, Laboratory for Molecular Biophysics, Physiology and Pharmacology, Univ. of Antwerp, Universiteitsplein 1, 2610 Antwerp, Belgium (e-mail: dirk.snyders{at}ua.ac.be)

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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