From the Department of Physiology and Biophysics and Rammelkamp Center for Research, MetroHealth Campus, Case Western Reserve University, Cleveland, Ohio 44109
The cytoplasmic half of S5 (5S5) has been identified as part of the inner mouth of the pore based
on evidence that mutations in this region greatly alter single channel conductance, 4-aminopyridine (4-AP) block
and the rate of channel closing upon repolarization (deactivation). The latter effect, suggestive of a role for 5
S5 in channel gating was investigated in the present study. The biophysical properties of chimeric channels, in which
the 5
S5 regions were exchanged between two host channels (Kv2.1 and Kv3.1) that differ in 4-AP sensitivity and
deactivation rate, were examined in a Xenopus oocyte expression system. Exchange of 5
S5 between Kv2.1 and
Kv3.1 confers steady-state voltage dependence of activation and rates of channel deactivation similar to those of
the donor channel. The involvement of voltage-dependent gating was confirmed by the observation that exchanging the 5
S5 segment of Kv2.1 with that of Kv3.1 confers a change from slow to fast deactivation kinetics by accelerating the decay of off-gating charge movement. We suggest that a conformational change that extends from the voltage-sensor in S4 to the region of the pore lined by S5 regulates the stability of the open state. Therefore, the
cytoplasmic end of S5, in addition to forming part of the conduction pathway near the inner mouth of the pore,
also participates in the conformational rearrangements associated with late steps in channel activation and early
steps in deactivation.
Voltage-gated K+ channels are integral membrane proteins that are assembled from four -subunits (MacKinnon, 1991
; Liman et al., 1992
), each of which contains
six transmembrane (S1-S6) segments (Tempel et al.,
1988
). Mutational analysis indicates that the ion permeation pathway contains elements of the S5-S6 linker
(Hartmann et al., 1991
; Yellen et al., 1991
; Yool and
Schwarz, 1991
; Holmgren et al., 1996
) and the cytoplasmic halves of S5 and S6 (Kirsch et al., 1993a
; Lopez et
al., 1994
; Shieh and Kirsch, 1994
). Side-chain substitutions at positions that are critical for determining permeation, however, have little effect on voltage-dependent gating currents (Taglialatela et al., 1992
). Instead,
attention has been focused on the S4 transmembrane
domain and its unique repeating motif of 5-7 positively
charged residues (Arg or Lys) each separated by two intervening hydrophobic residues. This motif is conserved in voltage-gated cationic channels including calcium and sodium channels (Catterall, 1988
) and is
thought to be part of the voltage sensor for channel
gating (Papazian et al., 1991
; Liman et al., 1991
; Logothetis et al., 1992
). Studies using cysteine mutagenesis, in combination with either chemical modification or
fluorescent labeling, indicate that the residues in the
S4 segment undergo a voltage-driven translocation
across the membrane electric field during activation
(Yang and Horn, 1995
; Mannuzzu et al., 1996
). The displacement of charged residues in the S4 contribute to
the gating currents (Aggarwal et al., 1996) that are responsible for regulating the voltage dependence of the
probability of opening. Charge movement kinetics are
directly related to the conformational rearrangements
of the channel protein associated with voltage-dependent gating transitions (Bezanilla and Stefani, 1994
)
and are very sensitive to hydrophobic substitutions in
the S4 and in other, downstream, regions of the channel (Zagotta and Aldrich, 1990
; Lopez et al., 1991
; McCormack et al., 1991
). Recently, it has been proposed
that in Shaker channels activation occurs via two voltage-dependent conformational rearrangements/subunit
that produce multiple transitional steps between closed
and open states (Bezanilla et al., 1994
; Sigg et al., 1994
;
Zagotta et al., 1994
). This behavior has been described
by models in which the early steps of activation are independent, whereas the later steps associated with the
final open
losed transitions involve concerted interaction of the subunits. In particular, both off-gating
charge movement and channel deactivation are limited
by a slow first closing transition that is not predicted by
independent movements of four identical subunits (Bezanilla et al., 1994
; Zagotta et al., 1994
). Although critical parts of the structural domains of the voltage sensor
and the ion conduction pathway have been identified,
the components responsible for rate-limiting transitions that couple movement of the voltage sensor to the channel opening are still unknown.
A clue to the coupling domain is that inhibition of
potassium currents by intracellular blockers such as
4-aminopyridine shows marked gating dependence
(Kirsch and Drewe, 1993; McCormack et al., 1994
; Yao
and Tseng, 1994
; Stephens et al., 1994
). In Shaker, Kv2.1 and Kv3.1, 4-AP preferentially enters and blocks
the activated channel. Once bound, the drug can be
trapped when the channel deactivates, such that its dissociation from the binding site requires channel reopening (Kirsch and Drewe, 1993
; McCormack et al.,
1994
). Furthermore, in Shaker channels it has been
shown that 4-AP interferes with a late step in activation
that leads directly to opening (McCormack et al., 1994
).
In view of the gating-dependent nature of 4-AP block,
the binding site and nearby residues may be located
within structural domains that undergo late conformational change during transitions between closed and
open states.
Previously, we have demonstrated that the cytoplasmic halves of both S5 and S6 specify differences in 4-AP
sensitivity between Kv2.1 and Kv3.1 (Kirsch et al.,
1993a; Shieh and Kirsch, 1994
). Furthermore, since
mutations in the S4-S5 linker and S5 have been shown
to affect gating kinetics (Zagotta and Aldrich, 1990
;
McCormack et al., 1991
; Kirsch et al., 1993a
; Holmgren
et al., 1996
), we have explored the possibility of a functional role for S5 in specifying differences in the kinetics of voltage-dependent gating between Kv2.1 and
Kv3.1. In the present report we show that chimeric
channels formed by reciprocal exchange of the cytoplasmic ends of S5 (5
S5) resulted in changes in the
rates of channel gating kinetics that were predictable
from the phenotypes of the host Kv2.1 and Kv3.1 channels. Our results suggest that the cytoplasmic end of
the S5 transmembrane segment regulates the rate of
the first closing transition.
Recombinant DNA and Mutagenesis
The host clones Kv2.1 (DRK1, Frech et al., 1989) and Kv3.1
(NGK2, Yokoyama et al., 1989
), and chimeric constructs were
propagated in the transcription-competent plasmid vector pBluescript SK(
) in the DH5
MCR competent cells (Gibco BRL,
Gaithersburg, MD). Details of the preparation of chimeras has
been described previously (Kirsch et al., 1993a
). Briefly, the
5
S5/Kv2.1 chimera had a peptide sequence identical to the host
Kv2.1 clone except that in the 5
half of the S5 segment, Leu327,
Gly328, and Leu332 (Kv2.1 numbering system) in the putative cytoplasmic end were replaced, respectively, with Phe, Leu, and Ile,
which correspond to the equivalent residues of the Kv3.1 donor.
5
S5/Kv3.1 is a chimera between Kv3.1 and Kv2.1 in which only the
Phe345, Leu346, and Ile350 (Kv3.1 numbering system) in the 5
end of
S5 of the host Kv3.1 clone were replaced by Leu, Gly, and Ile, which
correspond to the equivalent residues of the Kv2.1 donor.
RNA Transcription and Oocyte Injection
DNA constructs were linearized at the 3 ends by digestion with
NotI for runoff transcription. In vitro transcription with T7 RNA
polymerase was performed using the mMessage mMachine Kit (Ambion Inc., Austin, TX). The amount of cRNA synthesized
(20-100 µg) was quantified by the incorporation of trace
amounts of [32P]UTP in the synthesis mixture. The final cRNA
product was resuspended in 0.1 M KCl at a final concentration of
250 ng/µl and stored at
80°C. The integrity of the final product
and the absence of degraded RNA was determined by a denaturing formaldehyde 1% agarose gel stained with ethidium bromide. The cRNA was diluted to the desired concentrations (1-10
pg/nl for single channel or whole-cell recording; 250 pg/nl to
obtain saturated channel expression for gating current recording) immediately before oocyte injection. Stage V and VI Xenopus
oocytes were defolliculated by collagenase treatment (2 mg/ml
for 1.5 h) in a Ca-free buffer solution (in mM): 82.5, NaCl; 2.5, KCl; 1, MgCl2; 5, HEPES (+100 µg/ml gentamicin), pH 7.6. The
defolliculated oocytes were injected with 46 nl of cRNA solution
(in 0.1 M KCl) and incubated at 19°C in culture medium (in mM):
100, NaCl; 2, KCl; 1.8, CaCl2; 1, MgCl2, and 5, HEPES; 2.5, pyruvic acid (+100 µg/ml gentamicin), pH 7.6. Electrophysiological measurements were performed 2-6 d after cRNA injection.
Whole-cell Current Recording
Whole-cell currents were recorded in oocytes using a two-intracellular microelectrode voltage clamp as described previously (Drewe et al., 1994). Briefly, sharp-tipped agarose-cushion micropipettes (0.2-0.5 M
; Schreibmayer et al., 1994
) were used as
voltage-sensing and current-passing electrodes connected to a
commercial voltage-clamp amplifier (OC725C; Warner Instruments, Hamden, CT). Linear leakage and capacitative transient
currents were subtracted online using a P/4 subtraction routine.
K+ tail current relaxation from whole-cell recordings was fit to a
monoexponential function to obtain a deactivation time constant.
Single Channel Recording
Cell-attached patch recording was performed after manual removal of the vitelline envelope. Isotonic KCl bathing solution was
used to zero the resting potential, and the absence of resting membrane potential was verified by rupturing the membrane
patch at the end of each experiment to allow direct intracellular
potential measurement. Holding and test potentials applied to
the membrane patch during the experiment are reported as conventional intracellular potentials. Channels were activated by
rectangular test pulses from negative holding potentials. Current
records were low pass filtered at 1-2 kHz (3dB, four-pole Bessel
filter), then digitized at 5-10 kHz. Linear leakage and capacitative currents were subtracted digitally using the smoothed average of 10-20 null traces in which no channel openings could be
detected. Open and closed dwell time analysis was performed using idealized records (half-amplitude criterion; Transit analysis
program, Vandongen, 1996
). Dwell time histograms were fit to
monoexponential probability density functions using a maximum likelihood estimate. Events of <0.3-ms duration were excluded from fitting to avoid error introduced by the limited recording bandwidth (1 kHz). Bursts were identified by setting a
threshold for the maximum closed interval between events
within a burst according to the criterion of Colquhoun and Sakmann (1985)
. The number of channels present in a patch was determined by observing the maximum number of channels open
simultaneously at voltages where the probability of channel being
open was high. Where appropriate, data are expressed as mean ± SEM.
Gating Current Recording
Gating currents were recorded from the membranes of oocytes
expressing a high density of ion channels using patch-clamp
methods (Heinemann et al., 1992). Sylgard-coated macropatch
pipettes with diameter >10 µm were used to record from excised, inside-out membrane patches containing many channels.
To eliminate the ionic currents, intracellular permeant cations
(bath solution) were replaced by N-methyl-glucamine, and extracellular permeant cations (pipette solution) by tetraethylammonium. Membrane currents were evoked by rectangular test pulses
from a holding potential of
90 mV. The records were low-passed filtered at 2 kHz (
3dB, four-pole Bessel filter) and digitized at 50 kHz. The linear components of residual leakage and
capacitative currents were subtracted online using a
P/4 procedure using a subtraction holding potential,
100 mV. Gating charge
(Q) was obtained from the time integral of the gating current for
the duration of the test pulse. The charge vs. voltage (Q-V) curve
was generated by plotting normalized charge as a function of
voltage. Each data set was fitted by a Boltzmann equation:
![]() |
where V is the pulse potential, V0.5 is the half-activation potential, z is the effective valence, F is Faraday's constant, R is the universal gas constant, and T is the absolute temperature. The decay phase of gating current was fit to a biexponential decay function to determine the fast and slow components.
Numeric simulations of ionic and gating currents were obtained using Axovacs software (Axon Instruments, Foster City, CA) modified to accept Markov gating schemes (provided by Dr. Stephen Jones, Case Western Reserve University).
Solutions and Drugs
For gating current measurement, the oocyte was bathed in solution containing (mM): 120, N-methylglucamine; 120, glutamate; 10, EGTA; and 10, HEPES, pH 7.2 with N-methylglucamine. The pipette solution contained (mM): 122.5, tetraethylammonium hydroxide; 122.5, methane sulfonic acid; 2, CaCl2; 10, HEPES, pH 7.3 with NaOH. A modified Ringer's solution for whole-cell recording consisted of (in mM): 120, NaCl; 1, CaCl2; 2, MgCl2; 10, HEPES, pH 7.2 (with Tris-OH). The desired high external K+ solution was made by substituting the NaCl with KCl. Depolarizing isotonic KCl bath solution for single channel recording consisted of (in mM): 100, KCl; 10, EGTA; 10, HEPES, pH 7.3. Pipette solution was normal frog Ringer's solution containing (in mM): 120, NaCl; 2, KCl; 2, CaCl2; 10, HEPES, pH 7.2. When high K+ concentration was needed in the patch pipette, the Na+ was replaced with desired concentration of K+. Bathing solution flowed continuously at a rate of 3 ml/min. All electrophysiological measurements were made at room temperature (21-23°C).
Effects of S5 Chimeric Mutations on Macroscopic Ionic Currents
We examined the effects of reciprocal exchanges, between Kv2.1 and Kv3.1, of the cytoplasmic half of transmembrane segment, S5 (5S5), in chimeric channels.
The influence of 5
S5 on channel gating was readily observed at the macroscopic level in records obtained
from cell-attached membrane patches containing many
channels ("macropatches," Fig. 1). Inward tail currents
were evoked in patches exposed to elevated extracellular [K+] by a double pulse protocol in which a variable
test step (
120 to +40 mV) was applied immediately
after a fixed conditioning pulse to +60 mV that evoked
maximum activation. At test potentials more negative
than the activation range (
120 to
40 mV), the time
course of decay of the tail currents reflects the voltage-dependent closing rate (deactivation) of the channels.
The time constant of deactivation (Fig. 2) was estimated by fitting the tail currents during the test pulse
to a monoexponential decay. The fit excluded the first
0.1 ms of the test pulse to allow for clamp settling. As
shown in Fig. 2, the time constant was voltage dependent such that deactivation became progressively
slower at depolarized test potentials that approached
the activation range (>
40 mV). Chimeric substitutions of S5, however, markedly changed the rate of deactivation. The 5
S5/Kv2.1 chimera (Fig. 1 B) that contained the cytoplasmic half of S5 from the fast-gating
Kv3.1 channel (Fig. 1 C), showed markedly faster deactivation rates compared to its host Kv2.1 channel (Fig. 1
A). At strongly hyperpolarized test potentials (
150 to
60 mV; Fig. 2) where the forward rates of activation were negligible, the faster decay of the tail currents reflected primarily an acceleration of the channel closing
rate. In the reciprocal experiment, the 5
S5/Kv3.1 chimera (Fig. 1 D) that contained the cytoplasmic half of
S5 from the slow-gating Kv2.1 channel, showed a 10-50-fold slowing of deactivation throughout the test potential range compared to its host Kv3.1 channel (Fig. 1
C), consistent with participation of the 5
S5 region in
controlling the rate of channel closing.
In contrast, to the effects of S5 exchange on the deactivation rate, the effects on activation were much less
marked. Visual inspection of the rise time of outward
currents during conditioning steps to +60 mV (Fig. 1)
suggests that the activation kinetics of the chimeric
channels resembled that of the host channel. A quantitative comparison of the kinetics of the late phase of activation associated with the final transition to the open
state (Hoshi et al., 1994) was obtained by determining
activation time constants from monoexponential fits of
50-90% of the rising phase of the test pulse currents.
Thus at a test potential of +60 mV for Kv2.1 and the
5
S5/Kv2.1chimera, respectively, activation time constants of 11.5 ± 0.3 ms (n = 10) and 21.2 ± 0.2 ms (n = 9) were obtained; and for Kv3.1 the 5
S5/Kv3.1 chimera, respectively, 3.5 ± 0.2 ms (n = 5) and 2.7 ± 0.7 ms (n = 8) were obtained. It should be noted that the
chimeric mutations caused shifts in the steady-state voltage dependence of activation (see below) that
markedly affect activation time constants at test potentials that evoke less than maximum activation. However, the large differences between deactivation time
constants in the host and chimera for both sets of channels over an 80 mV range (Fig. 2) and the similarities
between the activation time constants argue that
changes in the channel deactivation rate by exchanging the 5
S5 region are not secondary to shifts in the
steady-state voltage dependence of activation. These results suggest that the 5
S5 region exerts rate-limiting
control of gating transitions primarily during the deactivation process.
Effects of S5 Mutations on Microscopic Gating
Since deactivation is a macroscopic property that can
be influenced by secondary transitions between closed
states, we examined the rate of channel closing directly
by single channel recording. Single channels (Fig. 3)
were activated by conditioning steps to +40 mV from a
holding potential of 60 mV, and the duration of the
open state was determined during the return to a test
potential of
80 mV. The patch pipette contained 60 mM K+ to facilitate the measurement of inward K+ current. In Kv3.1 (Fig. 3 A) rapid deactivation was correlated with extremely brief openings (mean open time = 1.6 ± 0.2 ms, n = 3 patches) during the
80-mV test
potential and no reopenings were observed. The ensemble average from 500 traces of single channel recordings gave a tail current with deactivation time constant of 3 ms obtained by fitting a single exponential
decay. This value is similar to the mean open time at
80 mV and to the deactivation time constant observed in whole-cell recordings of tail currents. By contrast, in Kv2.1 (Fig. 3 C) bursts of openings (mean burst
duration = 10.0 ± 2.2 ms, n = 6 patches) were recorded upon return to
80 mV. The burst duration
corresponded to the time constant of deactivation obtained from the ensemble average currents (
= 9.1 ± 1.7 ms, n = 6 patches). The fast and slow kinetics of
open to closed transitions in Kv3.1 and Kv2.1, respectively, were reciprocally transferred by swapping the cytoplasmic half of S5, as shown in 5
S5/Kv3.1 (Fig. 3 B)
and 5
S5/Kv2.1 (Fig. 3 D). Thus the 5
S5/Kv3.1 (Fig.
3B) mutation of Kv3.1 caused the mean open time at
80 mV to increase from 1.6 to 19.3 ms (n = 3 patches), and multiple reopenings were observed. Conversely, the 5
S5/Kv2.1 (Fig. 3 D) mutation of Kv2.1 resulted in channels with reduced conductance (Kirsch
et al., 1993b
) and marked changes in kinetics; during
depolarizing test pulses the channels opened briefly
(mean open time 0.9 ms ± 0.2, n = 4 patches) but repetitively. Unlike the host Kv2.1 channels, upon repolarization to
80 mV the /Kv2.1 channels stayed open
very briefly and rarely reopened.
Mutation Effects on First Latency Distributions
The distribution of the latency to first opening provides
insight into the kinetics of activation during transitions
from rest to open states. Upon reaching the open state,
if channels do not readily close into either an inactivated or closed state, the cumulative distribution of first
latencies will follow closely the time course of macroscopic activation (Hoshi et al., 1994). Alternatively, if
the open state is unstable such that transitions into
closed states are rapid relative to transitions from
closed to open states, the time course of first latencies
will be slower than that of macroscopic activation. Single channel recordings were made in the cell-attached
mode using patch pipettes containing normal frog
Ringer solution (Fig. 4 A) to maximize outward currents. Currents were evoked by test pulse potentials of
+40 mV from a holding potential of
60 mV. The
mean open time (pooled data, n = 3-4 patches) for
Kv2.1 and 5
S5/Kv2.1 was 12.0 ± 0.7 and 1.7 ± 0.2 ms,
respectively, and 16.5 ± 1.5 and 25.0 ± 2.8 ms for Kv3.1 and 5
S5/Kv3.1. In addition, comparison of the traces
obtained from Kv2.1 and 5
S5/Kv2.1 patches revealed a
mutation-induced decrease in the steady-state probability of opening (Po) in the 5
S5/Kv2.1 chimera due to a
simultaneous reduction in open time and an increase
in duration of the closed time intervals. At a test potential of 60 mV (data not shown) the average Po obtained from traces in which only one channel appeared to be
active was 0.91 and 0.05, respectively, for Kv2.1 and
5
S5/Kv2.1 channels (n = 3-4 patches). Moreover, the
time-to-first opening from the beginning of the test
pulse (first latency, Fig. 4 B) was relatively slow in the
chimera (open circles) compared with its host channel (filled circles). From the cumulative distributions of first
latencies we obtained an average t1/2 (half-rise time of
the cumulative distribution of latencies, n = 3-4
patches) of 1.5 ± 0.1 and 1.6 ± 0.1 ms for Kv3.1 and
the 5
S5/Kv3.1 chimera, respectively; and 19.3 ± 5.2 ms for Kv2.1 and 58.7 ± 7.9 ms for the 5
S5/Kv2.1 chimera. Since first latency measures the delay associated
with transit through multiple closed states before channel opening, our results indicate that chimeric mutations of S5 had no effect on the rate-limiting steps of activation when Kv3.1 was the host but caused at least a
twofold slowing when Kv2.1 was the host channel. It
should be noted that the first latency time course is
markedly affected by the correction of the data for the
number of channels in the patch (i.e., increasing the
number of channels results in apparently shorter first
latencies). This was not a problem for Kv2.1, Kv3.1, or
5
S5/Kv3.1 where the steady-state Po was near unity,
and single channel patches were easily obtained. However, for 5
S5/Kv2.1 channels, because of their extremely low Po, single channel patches could not be obtained, and the number of channels in each patch was
likely to be underestimated. Therefore, the effect of
this mutation on the corrected first latency distributions should be considered a minimum approximation
of the actual slowing.
Kinetic Model Simulation of S5 Mutations
The primary effect of the 5S5 exchange at the macroscopic level appears to be an alteration of the deactivation rate, and at the microscopic level, these effects
may be correlated with changes in transition rates associated with states near the open state of the fully activated channels. As a starting point we suggest that the
effects of 5
S5 mutations on Kv2.1 channels can be explained by a simple sequential scheme:
![]() |
We assume that transitions C1 C2 and C2
C3 are voltage dependent with forward and backward rate constants at 0 mV of 60 and 100 s
1, respectively; and effective valences (z) of 0.63 and
1.0, respectively. The
transition C3
O4 was assumed to be very rapid and
voltage insensitive, corresponding to the properties of
the mean closed interval within bursts at test potentials
in the range 0 to +60 mV (Taglialatela et al., 1993
). By
contrast, the reverse rate O4
C3 was assumed to be
slightly voltage dependent, corresponding to the properties of the mean open time in the range
90 to +80 mV (Fig. 5). A reasonable approximation to the measured single channel data was obtained by setting the
C3
O4 rate to 1,000 and 20 s
1 in Kv2.1 and 5
S5, respectively; and by setting the reverse rate, O4
C3 at 0 mV (z = 0.2 e
), to 150 and 1,000 s
1 in Kv2.1 and
5
S5/Kv2.1, respectively. The other voltage-dependent rate constants were assumed to be unaltered by the mutations, and no changes in effective valences were
made. In 5
S5/Kv2.1 channels these changes in rate
constants have the observed effect of markedly shortening the mean single channel open time from 7 to 1 ms,
lengthening the major closed interval from 1 to 50 ms
and markedly reducing open probability during maximal activation (from 0.86 to 0.04). As shown in the simulated whole cell current records (Fig. 6, A and B),
such a model predicts that in 5
S5/Kv2.1, tail current
time course will become very rapid compared to that of
the host channel and will approach the mean single
channel open time at test steps in the range
120 to
80 mV; i.e., the channel opens only once during the
tail, as observed in Fig. 3. Moreover in 5
S5/Kv2.1, because of its slow entry into O4 relative to its rapid exit
rate (O4
C3) the first latency time course (Fig. 4) will
be slow even though macroscopic activation (Fig. 6 A)
is relatively unaffected compared with Kv2.1.
The model makes several testable predictions (Fig. 6,
C and D): (a) the time course of the gating charge
movement upon repolarization from a test step that
maximally activates the channel should be much faster
in 5S5/Kv2.1 than in Kv2.1, whereas the time course during the activation step should be unchanged (Fig. 6
C); (b) the voltage dependence of the steady-state activation (Po-V relationship) should be shifted to more
positive potentials in 5
S5/Kv2.1 compared with Kv2.1
and will fail to saturate in the measurable range (i.e., below +80 mV, Fig. 6 D); (c) the steady-state voltage dependence of gating charge movement (Q-V relationship, not shown) will not change.
Effects of Mutation on Gating Currents
Gating currents were measured to verify the model and
to determine whether ion occupancy could be involved
in the observed kinetic changes. Deactivation, but not
activation, rates in delayed rectifier K+ channels are
thought to be sensitive to ion occupancy of the pore
(Swenson and Armstrong, 1981; Matteson and Swenson, 1986
). Therefore, we were concerned that the selective effects of S5 chimeric mutations on the closing
rate of the channels might be the indirect result of mutation-induced changes in ion occupancy. We addressed these issues by measuring gating currents in
the presence of impermeant ion substitutes. Gating
currents were recorded using macropatch methods
(Heinemann et al., 1992
) in Xenopus oocytes that expressed a high density of channels. Fig. 7 presents a
comparison of the time course of gating currents in oocytes expressing Kv3.1, Kv2.1, and 5
S5/Kv2.1 chimera
(the level of expression of the 5
S5/Kv3.1 chimera was
too low to allow resolution of the gating currents).
Each panel shows a gating current-voltage family evoked by test pulses (20-ms duration,
80 to +40 mV amplitude, Kv2.1 and 5
S5/Kv2.1 chimera; or
60 to +60 mV,
Kv3.1) in 20-mV increments from a holding potential
of either
100 mV (Kv2.1 and 5
S5/Kv2.1 chimera) or
90 mV (Kv3.1). Test pulses evoked outward, ON-gating current (Ion) that rose to a peak whose amplitude was
dependent on test pulse potential (Fig. 7 A). Ion then
decayed to the baseline in a biphasic manner. The relative contributions of the fast and slow components of
the decay were voltage sensitive: at low depolarizations (<+40 mV), the slow component of decay phase was
prominent, whereas at high depolarizations (>+40
mV) the fast component dominated. At the end of the
test pulse the return to the holding potential evoked inward OFF-gating current (Ioff) transients that decayed rapidly to the baseline.
In comparing the two host channels (Fig. 7) the most
striking feature of the gating currents was the marked
acceleration of the kinetics of both Ion and Ioff in Kv3.1
(A) compared with Kv2.1 (B). The chimeric 5S5/Kv2.1
channel (C) shows a mixture of the two host gating
phenotypes: the time course of Ion was similar to that of
the host Kv2.1 channel whereas the time course of Ioff resembles that of the donor Kv3.1 channel. Consistent
with the pattern observed in Kv3.1 (A), the decay of Ion
and Ioff in Kv2.1 and 5
S5/Kv2.1 exhibited both fast and
slow phases. A quantitative comparison between gating
current kinetics among Kv3.1, Kv2.1, and 5
S5/Kv2.1
was made by fitting exponential decay functions to the
gating currents. In each of the three channels, the decay of Ion could be accurately fit by to a biexponential
function in which the time constant of the slow phase
was about 10-fold slower and its amplitude 10-fold
smaller than those of the fast time phase. As plotted in
Fig. 8 A, at any given test potential Ion was dominated by
a fast time constant that was substantially faster in Kv3.1
than in Kv2.1. Likewise for Ioff (Fig. 8 B) the decay time
course observed in Kv3.1 was markedly faster in Kv3.1 than in Kv2.1. Substitution of the cytoplasmic end of S5
in Kv2.1 with that of Kv3.1 gave a hybrid kinetic pattern. Fig. 8 A shows that for Ion, the predominant time
constant observed in 5
S5/Kv2.1 was unchanged from
that of Kv2.1 and was much slower than that of Kv3.1.
Fig. 8 B shows that for Ioff, both the time course of decay for 5
S5/Kv2.1 channels was similar to that observed in Kv3.1 and faster than that seen in Kv2.1 channels. Thus, transplanting the cytoplasmic half of S5
from Kv3.1 into Kv2.1 converted the kinetics of Ioff to
resemble those of the donor channel, Kv3.1, but allowed retention of the on-gating kinetics of the host
Kv2.1 channel. These results, although obtained under
markedly different ionic conditions, closely resemble
the effects on ionic currents.
Effects of S5 Mutations on Steady-state Voltage Dependence of Activation
The amount of charge displaced during activation was
obtained from the time integral of Ion at each test potential for the host channels Kv2.1 and Kv3.1 and for
the chimeric 5S5/Kv2.1 channel in which Kv2.1 was
the host and Kv3.1 the donor channel. The gating
charge activation curve (Q-V curve) was generated by plotting the normalized charge displaced versus test
potential, and the curve was fitted with single Boltzmann equation (Fig. 9 A). The steady-state activation
curve (G-V curve, Fig. 9 B) was obtained by plotting
normalized ionic conductance versus test potential in
separate experiments. The fitted half-activation potential (V0.5) of the Q-V relationship was 13 ± 2 mV and
25 ± 4 mV for Kv3.1 and Kv2.1, respectively. Similarly, the half-activation potentials of the G-V relationship were 20 ± 2 and 10 ± 1 mV for Kv3.1 and Kv2.1,
respectively. The effective valence for Kv3.1 was 2.8 ± 0.2 and 2.0 ± 0.3 e
for Kv2.1. In the 5
S5/Kv2.1 chimeric channel, the Q-V curve remained the same as
that of Kv2.1, with V0.5 of
22 ± 3 mV and effective valence of 1.7 ± 0.1 e. However, the 5
S5/Kv2.1 substitution caused a positive shift in the G-V curve with apparent V0.5 of 27 ± 4 mV compared with 10 ± 1 mV in
Kv2.1. The actual shift may have been larger than indicated by curve-fitting of normalized data since the
5
S5/Kv2.1 G-V curve did not saturate at the most positive test potentials available experimentally. These results are comparable to those predicted by model simulation (Fig. 6 D). In the model for the O4
C3 step is
slightly voltage dependent such that its rate constant
becomes slower with depolarization and should contribute to the voltage dependence of Po. In Kv2.1, the
effect is negligible since this rate is quite slow compared to the voltage independent C3
O4 step, and Po
saturates at a level of about 0.9 at test potentials >0 mV.
In contrast, for the 5
S5/Kv2.1 chimera, the O4
C3
step dominates the final step in activation and is responsible for the very low Po observed experimentally.
The voltage dependence of this step causes Po to continue creep upwards at test potentials >+40 mV.
For 5S5/Kv3.1 the midpoint of the G -V curve was
25 ± 2 mV consistent with a transfer of at least part of
the gating phenotype of the donor Kv2.1 channel.
Thus, a reciprocal change in the steady-state voltage dependence activation was produced in Kv2.1 or Kv3.1 by swapping the cytoplasmic half of S5 transmembrane
segment.
Structural Basis of Gating Differences between Kv2.1 and Kv3.1
Kv2.1 and Kv3.1 are related voltage-gated K+ channels that have distinct gating phenotypes: Kv2.1 has markedly slower activation and deactivation rates compared with Kv3.1, and operates over a more negative voltage range. The effect of exchanging the cytoplasmic halves of S5 between the fast-gating Kv3.1 and slow-gating Kv2.1 channels had predictable effects on the rates of macroscopic deactivation in the chimeras. The macroscopic activation rates were relatively unaffected. At the microscopic level this suggests that S5 contributes to the regulation of the final step in activation that leads directly to channel opening and, upon repolarization, is the first closing step in deactivation.
A simple kinetic explanation for most of our results is
that in the rapidly deactivating 5S5/Kv2.1 chimera the
open state is destabilized by marked acceleration of the
closing rate and slowing of the opening rate. This results in decreased probability of opening and a rightward shift in the G -V curve along the voltage axis with
little change in Q-V, and a marked decrease in the time constants of both deactivation and Ioff. Conversely, the
electrophysiological phenotype of the 5
S5/Kv3.1 chimera was consistent with stabilization of the open state.
Our kinetic model is inaccurate, however, in its prediction that the time course of activation should be unchanged in 5S5/Kv2.1 channels compared with Kv2.1
(Fig. 6), whereas an almost twofold slowing of the late
phase of activation was observed in the mutant channels (Fig. 1). A more accurate model could be constructed based on kinetic analyses of gating in Shaker K
channels which indicate that during activation the
channel undergoes a series of voltage-dependent transitional steps from a resting state to a pre-open, permissive state, followed by a concerted movement that
opens the ion conduction pathway (Bezanilla et al.,
1994
; Sigg et al., 1994
; Zagotta et al., 1994
). The concerted nature of the final step may involve functional
interactions between subunits such as those observed
previously to originate from the structural components
of the pore itself (Kirsch et al., 1993b
; Ogielska et al.,
1995
). Therefore, changes in an additional free parameter corresponding to the allosteric interaction could
be invoked to account for the slowing of macroscopic
activation in 5
S5. However, this does not alter our
main conclusion that the 5
S5 region regulates the final step.
Our gating current results suggest that fundamental
differences in the kinetics of the voltage-dependent
charge movements underlie the observed differences
in the kinetics of the ionic currents between the two
host channels. Thus, we found that the decay of both
Ion and Ioff in Kv3.1 was significantly faster in Kv3.1 compared with Kv2.1. Also, we found that both the Q-V and the G -V relationships were shifted in the positive direction, and the steepness of the curves was greater in
Kv2.1 compared with Kv3.1. However, the 5S5/Kv2.1
chimera retained the Q-V curve and Ion kinetics of the
host channel. Therefore, the differences between Kv3.1
and Kv2.1 cannot be attributed entirely to different 5
S5 residues but may reflect the contribution of other
regions of the protein. Differences in the number of
charged residues in S4 are an obvious candidate; the
first Arg residue in the S4 segment of Kv3.1 (corresponding to Arg362 in Shaker) is replaced by an uncharged Gln in Kv2.1. However, in Shaker (Pappazian,
1991) and its mammalian homolog (Kv1.1; Liman et
al., 1991
; Logothetis et al., 1992
), Gln substitution shifts
the G-V curve in the wrong direction (i.e., to more positive potentials). Alternatively, multiple substitutions
among nonconserved, uncharged residues in the S4
can have marked effects on the position of the G-V
curve along the voltage axis (Logothetis et al., 1993
). Whether mutations of these nonconserved residues can
affect the position of the Q-V curve or the kinetics of
the gating currents has not been determined. However,
Val substitution for highly conserved Leu residues in
Shaker S4 segments is known to have dramatic effects on
the position of both the G-V and Q-V curves (Schoppa
et al., 1992
).
In the present study, each chimeric construct included multiple mutations in the cytoplasmic half of S5
in Kv2.1 (Leu327 Phe, Gly328
Ile, Leu332
Ile) or Kv3.1
(Phe345
Leu, Ile346
Gly, Leu350
Ile). We found previously (Shieh and Kirsch, 1994
) that in Kv2.1 the point
mutations, G328I or L332I were more effective than
L327F in shifting the G-V curve to the right and in reducing single channel open time. L332I, in particular,
reduced the single channel open probability from
about 0.85 to 0.05, a value similar to that obtained in
the 5
S5/Kv2.1 mutant. Moreover, in Shaker, Phe401, the
corresponding residue, when mutated to Ile also shifts
the G-V curve to more positive potentials (Zagotta and
Aldrich, 1990
). The highly conservative nature of the
Kv2.1 Leu/Ile substitution and the similarity to Shaker
results suggests that this position may be a critical determinant of differences in gating between channel isoforms. Previous work in Shaker also has shown that point mutations involving Leu residues in S5 can affect
the rate and apparent voltage dependence of channel
activation (McCormack et al., 1991
). For instance, the
first Leu in S5 of Kv2.1 (corresponding to Leu496 in
Shaker) is part of a highly conserved leucine-heptad repeat structure that has been suggested as a participant
in allosteric interactions between S4 and the pore (McCormack et al., 1991
; McCormack et al. 1994
). Leu/Val
substitution in S5 (Leu396 in Shaker) caused a moderate
negative shift in the G-V curve, but in Kv2.1, Phe substitution of the corresponding Leu (Kv2.1 L327F) had little effect on gating (Shieh and Kirsch, 1994
). Whether other strictly conserved residues such as Leu329, Leu330
or Phe333 are functionally critical remains to be determined.
In summary, our results indicate that residues in 5S5
regulate channel deactivation. The effects of mutation
on tail current kinetics seem to be specific since the
mutual exchange of three residues between Kv2.1 and
Kv3.1 produced reciprocal effects on channel closing
rate. The change in deactivation kinetics was accompanied by a change in the kinetics of off-gating charge
movement. The results suggest that the 5
S5, in addition to forming part of the inner mouth of the pore
(Kirsch et al., 1993a; Shieh and Kirsch, 1994
), helps to
regulate the final transition to and from the open state.
Original version received 27 January 1997 and accepted version received 8 April 1997.
Address correspondence to Dr. Glenn E. Kirsch, Rammelkamp Center for Research, R327, MetroHealth Medical Center, 2500 MetroHealth Dr., Cleveland, OH 44109. Fax: 216-778-8282; E-mail: gek3{at}po.cwru.edu
A brief account of these results has been reported previously (Shieh, C.C., K.J. Greene, and G.E. Kirsch. 1996. Biophys. J. 70:A144).We thank W.-Q. Dong and C.-D. Zuo for expert oocyte injection and culture. We also thank Dr. Stephen W. Jones for programs and advice on kinetic modeling.
This work was supported by National Institutes of Health grant NS29473 to G.E. Kirsch and American Heart Association Grant-in-Aid, Northeast Ohio Affiliate to C.C. Shieh.