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Biophys J, April 2000, p. 1852-1861, Vol. 78, No. 4
School of Biomedical Sciences, University of Leeds, Leeds LS2 9JT, England
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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The positively charged S4 region of voltage-dependent potassium channels moves outward during depolarization, leading to channel opening, but possible movement of the negatively charged S2 region may be more complex. Here we have studied possible movement of the S2 region of the slowly activating human voltage-dependent potassium channel hKv2.1. For this, cysteine mutants in the S2 region were expressed in Xenopus oocytes by injection of cRNA. Whole-cell currents were measured using the two-electrode voltage-clamp technique, and the effect of the membrane-impermeable cysteine-binding reagent parachloromercuribenzenesulfonate (PCMBS) was studied. For mutant S223C (located just outside the membrane in the S2 region), PCMBS inhibited currents and caused faster deactivation of tail currents. The time course of reactivity of PCMBS on tail current amplitudes was faster at more negative holding potentials. There was no effect of PCMBS on potassium channel currents for mutants D225C, N226C, A230C, and V232C. These data suggest that residue S223 is exposed to the extracellular phase at normal resting potentials, making it accessible to PCMBS, but upon depolarization there is a conformational change, making it less accessible, possibly by a local rather than global movement of S2 residues into the membrane. Voltage-dependent movements of nearby residues could also explain the results.
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Voltage-gated potassium channels play an
important role in controlling electrical activity and proper
functioning of excitable cells. Numerous voltage-gated
outward-rectifying potassium channels have now been cloned and share a
common structure comprising four 
-subunits surrounding a central
aqueous pore, with each -subunit consisting of six hydrophobic
transmembrane segments, S1-S6 (reviewed by Armstrong and Hille, 1998
;
Pongs, 1992). The human hKv2.1 (DHK1 or hDRK1) channel belongs to this
family of voltage-gated outward rectifier potassium channels and is
characterized by slowly activating currents, noninactivating currents
when expressed in Xenopus oocytes (Albrecht et al., 1993).
The roles played by some of the transmembrane domains have been
extensively studied. The S4 segment consists of conserved basic
residues at every third position, and detailed work has shown that this
segment plays an important role in voltage sensing by the channel
(Liman et al., 1991; Papazian et al., 1991; Logothetis et al., 1992;
Perozo et al., 1993; Shao and Papazian, 1993; Aggarwal and MacKinnon,
1996). The S2 and S3 segments contain negatively charged residues, and
more recent work has also begun to implicate these regions in voltage
sensing (Papazian et al., 1995; Planells-Cases et al., 1995; Seoh et
al., 1996; Tiwari-Wood-ruff et al., 1997). The linker between the
S5-S6 segments (H5 or P region) contains a highly conserved sequence
that is invaginated into the membrane from the extracellular face and
lines the outer mouth of the pore, forming the selectivity filter
(MacKinnon and Yellen, 1990; Hartmann et al., 1991; Yellen et al.,
1991; Yool and Schwartz, 1991; Doyle et al., 1998). The S5 and S6
regions line the part of the pore internal to the selectivity filter
(Choi et al., 1993; Kirsch et al., 1993; Lopez et al., 1994; Holmgren
et al., 1997).
More recently, studies of conformational changes in channels have been
made using cysteine-scanning mutagenesis. More specifically, movement
of the S4 region during depolarization has been demonstrated by the
application to Shaker potassium channels of
membrane-impermeable cysteine-binding reagents (Larsson et al., 1996;
Yusaf et al., 1996; Baker et al., 1998), thus probing the accessibility
of cysteine-mutated residues. In this way it was possible to show that,
upon depolarization, the S4 segment appears to move outward by around
seven amino acids. Complementary to this approach has been the use of
fluorescent cysteine-binding probes, which provide useful information
on changes in the local environment of S4 during depolarization
(Mannuzzu et al., 1996). Strong electrostatic interactions exist
between the negatively charged residues in S2/S3 and the positive
residues in S4; these interactions are important for folding and
functioning of the channel (Papazian et al., 1995; Tiwari-Woodruff et
al., 1997). It seems plausible to hypothesize that as the S4
transmembrane segment moves outward upon depolarization, the S2
segment, with its negatively charged residues, might move inward.
Studies have been made of the changes in the local environment of the
S2 region of the Shaker channel, using fluorescent probes
(Cha and Bezanilla, 1997), and local rather than global movements of
the S2 region have been suggested.
To investigate possible movement of the S2 region in this paper, we
have used cysteine-binding reagents to study the accessibility of
residues in the S2 region under varying degrees of depolarization. For
this we have used cysteine mutants of the human
K+ channel clone hKv2.1, and we have tested
accessibility using the membrane-impermeable sulfhydryl-reactive
reagent parachloromercuribenzesulfonate (PCMBS) in voltage-clamped
oocytes. This reagent has already proved useful in inhibiting currents
in the S4 region (Yusaf et al., 1996).
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MATERIALS AND METHODS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Preparation of DNA
The hKv2.1 clone in pGEM-He-Juel (provided by O. Pongs, Hamburg,
Germany) was transformed into Escherichia coli DH5
competent cells (Sambrook et al., 1989). DNA was isolated with a
miniprep kit (Promega) according to the manufacturer's instructions.
Preparation of mutants
The mutants S223C, D225C, N226C, Q228C, A230C, and V232C were
produced by polymerase chain reaction (PCR) methods (Innis et al.,
1990). PCR reactions were carried out (Sambrook et al., 1989) using Pfu
polymerase enzyme for 30 cycles (40 s at 95°C, 1 min at 60-66°C,
and 2 min at 72°C).
First-round PCR reactions were carried out using a forward primer,
5'-ATGCCGGCGGGCATGACGAA-3', and a reverse mutagenic primer (incorporating the desired base change), as well as a forward mutagenic
primer and a reverse primer, 5'-AACAATTTTCCCCAGGAGAGTCTTG-3'. Second-round PCR extension was carried out using the above products and
the above forward and reverse primers, to produce a ~1-kb product.
This latter product was then digested with the restriction enzymes
PpuMI and ApaI to produce a 250-bp fragment. The
wild-type hKv2.1/pGEM-He-Juel was also digested with the same enzymes,
and the mutagenic 250-bp fragment was ligated using
T4 ligase. The induced mutations were verified by
dideoxy sequencing (Sambrook et al., 1989).
To make RNA, the cDNAs for hKv2.1 were linearized with NotI, and capped cRNA was transcribed using T7 polymerase via a MEGAscript kit (Ambion). The RNA concentration was estimated using a Tris-acetate EDTA agarose gel and comparing with standard markers.
Electrophysiology
Xenopus oocytes (Dumont stage V or VI) were prepared
and injected with cRNA, and electrophysiological recordings were made as previously described (Wilson et al., 1994). Briefly, oocytes were
injected with 50 nl (containing 0.1-2.0 ng cRNA) of wild-type or
mutant cRNA. Oocytes were then incubated, at 19.7°C, for 24-48 h, in
multiwell tissue culture plates (one oocyte per well) containing modified Barth's solution (88 mM NaCl, 1 mM KCl, 2.4 mM
NaHCO3, 0.82 mM MgSO4, 0.33 mM Ca(NO3)2, 0.4 mM
CaCl2, 7.5 mM Tris-HCl, pH 7.6, 10,000 U/L
penicillin, 100 mg/L streptomycin). To record expressed membrane
currents from oocytes, the oocytes were held in a recording chamber (50 µl volume) and continually perfused (2 ml/min) at room temperature
(normally 22-25°C) with Ringer's solution (115 mM NaCl, 2 mM KCl,
1.8 mM CaCl2, 10 mM HEPES, adjusted to pH 7.2 with NaOH). Membrane currents were recorded by the two-electrode voltage-clamp technique with a Geneclamp 500 amplifier (Axon
Instruments). The amplifier was controlled via a CED 1401 Plus
interface and CED software. The current signal was sampled at 4 kHz and
filtered at 2 kHz. Electrodes were filled with 3 M KCl (0.5-2.0 M
resistance). To construct current-voltage relationships, the membrane
potential was held at 
80 mV, and 500-ms duration test depolarizations
(at 0.1 Hz) were applied in 10-mV increments, from 70 mV to +70 mV, and peak current amplitudes were measured. Twenty 10-mV hyperpolarizing steps (500 ms, 0.5 Hz) were applied and used to remove leak and capacitance currents. PCMBS was applied by continuous perfusion during
repetitive step depolarizations (500 ms, 0.1 Hz, +40 mV) from the
selected holding potential (usually 80 mV). To study tail currents,
oocytes were perfused with high potassium solution (100 mM KCl, 2 mM
MgCl2, 1.0 mM CaCl2, 10 mM
HEPES, pH 7.2), and test pulses were applied at 0.1 Hz. The membrane
potential was held at the selected holding potential (usually 80 mV),
and 100 ms prepulses to +40 mV were applied, followed by 350-ms test
pulses in 10-mV increments from 0 mV to 110 mV. Tail-current
amplitudes were measured 2 ms after the end of the prepulse, and the
deactivation time course was fitted to a single exponential.
Exponential fits to the deactivation time course were also used to
determine tail-current amplitudes by extrapolation back to the end of
the prepulse (particularly where PCMBS affected the time course); this
method gave results similar to those obtained by measurement of the
current amplitude at 2 ms (as expected, because deactivation time
constants were much longer than this), so the latter method was
generally used. PCMBS was applied at different holding potentials
during repetitive stimulation, the time constant of reactivity of PCMBS
was obtained by fitting current amplitudes to a single-exponential
curve for each cell, and mean values were obtained.
Many of the results for the mutants tested gave no effect of PCMBS, and the data for these are described but generally are not shown in this paper, in the interest of brevity and clarity. Data are expressed as the mean ± standard error of the mean (SEM), and Student's t-test was used to test statistical significance.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Characterization of wild-type and mutant currents
The membrane topology of hKv2.1 is illustrated in Fig. 1, which also shows the residues that have been mutated into cysteines in the S2 domain and in the S1/S2 linker. The wild-type or mutant cRNA was injected into Xenopus oocytes, and voltage-clamp recordings were made after 24-48 h.
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Characteristics of the wild-type channel are shown in Fig.
2. Mean current-voltage (I-V)
curves, shown in Fig. 2 A, were obtained during step
depolarizations from 80 mV, and the voltage dependence expected for
this outwardly-rectifying channel can be seen (Albrecht et al., 1993).
Fig. 2 B shows example recordings and illustrates the slow
activation time course and the minor inactivation characteristic of the
hKv2.1 channel. Wild-type tail currents were recorded in high potassium
solution (see Materials and Methods) during steps after a +40-mV,
100-ms prepulse; an example recording is shown in Fig. 2 D.
The mean tail-current I-V curve is shown in Fig. 2
C, reversing at around 5 mV as expected; mean deactivation time constants are plotted against voltage in Fig. 2 E.
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All the mutants studied produced functional channels when expressed in
oocytes, except for Q228C, which did not display detectable currents.
The I-V curves for the mutants S223C (Fig. 4 B),
D225C, N226C, A230C, and V232C (data not shown) were similar to the
wild-type I-V curves. The time courses (typical examples
shown in Fig. 3) of the mutant currents
were generally similar to wild type with slow activation and little
inactivation, except for N226C, which displayed an obvious inactivation
component. For mutant tail currents, I-V curves were again
generally similar to wild type (Fig. 5 B for S223C), as was
the deactivation time course of the tail currents. However, for mutant
D225C, the deactivation time constant (84.1 ± 12.4 ms at 70 mV
and 74.5 ± 4.5 ms at 30 mV) was voltage-independent (cf. Fig. 2
E for wild type).
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Effects of PCMBS on wild-type and mutant currents
The effect of PCMBS (100 µM) was tested using continuous
perfusion and repetitive stimulation every 10 s to +40 mV from a holding potential of 80 mV (Fig. 3). There was no effect of PCMBS on
wild-type and mutant channels, except for S223C, where there was a
partial inhibition by ~20% (p < 0.05). Study of
I-V curves before and after the application of PCMBS under
the same conditions showed similar features: no effect of PCMBS for
wild type (Fig. 2 A) and mutants, except for a small
reduction for mutant S223C (Fig. 4
B).
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Repetitive stimulation per se was not required for the inhibitory
action of PCMBS. This can be seen from Fig. 4 A, where
oocytes were held at 80 mV in the absence of pulsing for 2 min (a
time interval sufficient to reach a plateau value with pulsing; see above); the inhibition was still observed. Thus channel opening itself
is not required for PCMBS action, and indeed channel blocking would not
be expected consequent to binding to the S2 region, away from the pore.
As shown in Fig. 4, A and C, the inhibition by PCMBS was not affected by washing, as expected for covalent modification. Furthermore, the inhibition was completely reversed (Fig. 4 C) by the application of dithiothreitol (DTT), a reducing agent that removes PCMBS from cysteines (but which had no action on its own on S223C). This therefore confirms that PCMBS is indeed acting via cysteine modification rather than by some other nonspecific action.
The effects of PCMBS on tail currents for wild-type and mutant channels
were also studied. There was no effect of PCMBS applied at 80-mV
holding potential on tail-current I-V curves for wild-type channels (Fig. 2, C and D) and for all of the
mutants studied (data not shown), except for S223C, where an inhibition
of tail-current amplitude was seen (Fig.
5 B).
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The voltage and time dependence of the effect of PCMBS on tail-current
amplitudes was studied for S223C by applying the reagent at different
holding potentials during repetitive pulsing (to 100 mV from a
prepulse of +40 mV) (Fig. 5 A). As compared with wild type,
where no effect of PCMBS was observed, tail-current amplitudes were
always inhibited for S223C. The time course of reactivity of PCMBS on
tail-current amplitude was slower at depolarized holding potentials
(40 and 60 mV) as compared with more negative potentials (80 and
120 mV) (Fig. 5, A and D). The magnitude of the
effect on amplitude was variable, but because experimental scatter was
large, a possible voltage-dependent effect on amplitude was not
investigated further. There was no effect of PCMBS on deactivation time
constant for wild-type currents (Fig. 2 E) or for any of the
mutants, except again for S223C, where PCMBS caused faster deactivation
(Fig. 5, inset). Significant effects on the deactivation
time constant were observed when PCMBS was applied at intermediate
potentials (Fig. 5 C) rather than at either hyperpolarized or depolarized potentials.
In summary, the results in this section show that PCMBS was without effect on wild-type and mutant currents, except for mutant S223C, where inhibitory effects were observed. For the latter mutant, voltage-dependent effects of PCMBS (summarized in Fig. 5, C and D) were observed for deactivation time constants and for the time course of reactivity of PCMBS on tail-current amplitude. Channel opening per se was not required for PCMBS action, while reversal of the effect by DTT confirmed a specific action via binding to cysteine at 223.
Effects of other cysteine-binding reagents
The effects of PCMBS described above were not large, so we have investigated whether other cysteine-binding reagents (methanethiosulfonate ethyltrimethylammonium, MTSET, and thimerosal) had a greater effect and so might be easier to characterize. As for PCMBS, these reagents had no effect on wild-type currents (Fig. 6, A and C). For mutant S223C, MTSET produced an inhibition of ~12% (p < 0.05), and this was also reflected in the inhibition of the I-V curve in a voltage-independent way (Fig. 6 B). For thimerosal applied to this mutant, there was a small increase in currents (by ~10%, p < 0.05) with a corresponding small increase in the I-V curve, again voltage-independent (Fig. 6 D).
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Thus the other cysteine-binding reagents tested (MTSET and thimerosal)
had no greater effect than PCMBS and were therefore not examined
further. MTSET produced an inhibitory affect, as has generally been
observed elsewhere (Larsson et al., 1996), while thimerosal produced a
small increase, as has also been seen for other channels (Yao et al.,
1997).
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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In the experiments reported here, mutants S223C, D225C, N226C, Q228C, A230C, and V232C of the human outward-rectifier hKv2.1 were studied. Mutant Q228C did not express detectable currents in oocytes, while the other mutants displayed I-V curves similar to those of wild-type channels, although some of the mutants displayed altered time courses. Mutation to cysteine of some of the residues produced kinetic changes; the time courses of inactivation and deactivation were altered for mutants N226C and D225C, respectively. These residues may have roles in channel function.
The membrane-impermeable cysteine-binding reagent PCMBS was without effect on wild-type and mutant currents, except for mutant S223C, where there was a partial inhibition of currents. Furthermore, other cysteine-binding reagents tested (MTSET and thimerosal) had effects on mutant S223C but not on wild type. These results, together with the reversal of the effect of PCMBS by DTT, show that the cysteine at residue 223 specifically binds these reagents and that this residue is accessible and hence exposed from the extracellular side.
There were voltage-dependent effects of PCMBS when it was applied to mutant S223C, which were studied in detail for tail currents and are summarized in Fig. 5. The time course of reactivity of PCMBS on tail current amplitudes was faster at more negative potentials. These voltage-dependent results for the S223C mutant suggest that this residue is more accessible to the extracellular phase at hyperpolarized potentials. At very negative potentials, the deactivation time constant was less affected by PCMBS, but possibly this could be the result of a secondary effect rather than a decrease in accessibility, because the best measure of accessibility is the time of reactivity with PCMBS.
The decrease in accessibility at depolarized potentials can be
explained by an expected movement of the negatively charged S2 region
into the membrane when the cell is depolarized. A possible scenario for
this could be local movement of residue S223 rather than movement as a
whole of the S2 region. However, changes in accessibility could also
come about, for instance, by some local occluding conformational change
of adjacent regions/residues of the protein, so exposing or occluding
S223 during potential changes. So for instance, the effect could arise
as a side effect of possible movement of the S3-S4 linker, as in the
helical screw model (Durell and Guy, 1992). However, such proposed
movement of the S3-S4 linker may in fact not occur, and it could remain
fairly stationary (Aggarwal and MacKinnon, 1996, Mathur et al., 1997).
Our results showing a lack of effect of PCMBS on the other residues suggest either that amino acids D225, N226, A230, and V232 are not accessible to PCMBS or that binding of the reagent does not affect function. A reasonable suggestion is that residues 230 and 232 are not exposed to the extracellular environment, consonant with the proposed intramembrane (i.e., inaccessible) location of these residues. For residues 225 and 226, either they are not critical for function (indicating a local effect of PCMBS on residue 223), or they are not accessible to PCMBS (possibly because residues 225 and 226 are closer to the membrane than 223, and so might be obstructed by neighboring structures).
The effects of PCMBS on mutant S223C were small compared with the much
larger effects seen in studies of the S4 region with this reagent
(Yusaf et al., 1996). This partial inhibition of the S223C mutant
channel might suggest that only a small local conformational change at
this residue occurs when the membrane potential is changed; in any
case, binding of PCMBS does not seem to have induced major structural
changes causing a block of channel function. Absence of use-dependent
block suggests that PCMBS binding to the S2 region does not cause open
channel block, consonant with the location of S2 away from the pore.
It is interesting that deactivation was speeded up rather than slowed
by PCMBS. Possibly interference with the S2 region by PCMBS
paradoxically may allow the S4 region to move back into its resting
position faster after hyperpolarization, thus closing the channel
faster. This may be relevant to the suggestion that S2 gating charge
movement occurs before S4 movement (Seoh et al., 1996; Cha and
Bezanilla, 1997) for channel opening and presumably vice versa for
channel closing during deactivation.
Our previous work using PCMBS has shown that several contiguous amino
acids in the S4 region together become accessible upon depolarization,
consistent with a model (among others) of contiguous movement of
several amino acids out of the membrane (Yusaf et al., 1996). This
contrasts with the above results, where effects of PCMBS were seen
locally on residue S223 but not on the nearby extracellular residues
D225 and N226, which also might be expected to move inward toward the
S2 region. Previous work using fluorescent labeling of extracellular S2
cysteine mutants for the Shaker channel (Cha and Bezanilla,
1997) has also indicated local but not global movement of the S2
residues; significant changes in fluorescence with membrane potential
were found for two residues but not for two other residues nearby. Cha
and Bezanilla (1997) found, as we have, that the main effect of
cysteine probes was observed at intermediate potentials. Detailed
comparison between equivalent residues in Shaker and hKv2.1
suggests that Shaker T276 corresponds to hKv2.1 N226, and
yet there was a voltage-dependent change in fluorescence for T276 in
the study of Cha and Bezanilla but no effect for PCMBS in our study for
N226. Furthermore, Shaker P273 showed no fluorescence
changes with potential, while the corresponding residue S223 in hKv2.1
showed voltage-dependent effects. There are several possible
explanations for the differences. First, the fluorescent labels sense
localized changes in positions of nearby residues, whereas the studies
using PCMBS performed here determine accessibility. Clearly the
movement of surrounding residues could be such that surrounding
residues might move, so affecting fluorescence but without affecting
accessibility of PCMBS, and vice versa. Second, the nature of the
cysteine-binding probe can affect results; for instance, fluorescent
probes with differing charge can produce differing effects at the same
residue (Cha and Bezanilla, 1997). Third, differences between different
channel clones may be more apparent than real because alignments are
uncertain for the nonconserved loop regions; indeed the S1/S2 linker is 12 residues shorter for hKv2.1 than for Shaker.
In summary, our results suggest that some of the residues located extracellularly to the S2 region play some role in channel gating. In particular, residue S223 is accessible to PCMBS at normal resting potentials and therefore must be exposed to the extracellular phase. Nearby residues were not available to PCMBS. Upon depolarization, residue 223 becomes less accessible, probably as a result of a local inward movement rather than a simple movement en bloc of the negatively charged S2 region into the membrane.
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ACKNOWLEDGMENTS |
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We thank the Wellcome Trust for support and Prof. O. Pongs for the hKv2.1 clone and mutants A230C and N226C.
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FOOTNOTES |
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Received for publication 11 February 1999 and in final form 13 December 1999.
Address reprint requests to Prof. D. Wray, Biomedical Sciences, University of Leeds, Leeds LS2 9JT, England. Tel.: 44-113-233-4320; E-mail: d.wray{at}leeds.ac.uk.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
Biophys J, April 2000, p. 1852-1861, Vol. 78, No. 4
© 2000 by the Biophysical Society 0006-3495/00/04/1852/10 $2.00
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) and after (
) the
application of PCMBS (100 µM), using the repetitive stimulation
protocol (see Materials and Methods). Currents were normalized with
respect to the control current amplitude obtained in the absence of
PCMBS at a test potential of +70 mV (control current amplitude
12.3 ± 0.1 µA, n = 4). (B)
Current traces recorded for steps from 
deact)
before (
) was
significantly reduced by MTSET (p < 0.05) for
t > 2 min as compared to wild type and to initial
current values. MTSET had no significant effect on the current
amplitude for the wild-type channel (
). (B)
Current-voltage relationship curves before (