Contribution of the NH2
terminus of Kv2.1 to channel activation
Juan M.
Pascual1,
Char-Chang
Shieh2,
Glenn E.
Kirsch2, and
Arthur M.
Brown2
1 Center for Molecular
Recognition, Columbia University, New York, New York
10032; Department of Molecular Physiology and Biophysics, Baylor
College of Medicine, Houston, Texas 77030; and
2 Department of Physiology and
Biophysics and Rammelkamp Center, MetroHealth Campus, Case Western
Reserve University, Cleveland, Ohio 44109
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ABSTRACT |
Opening and closing of voltage-operated channels requires the
interaction of diverse structural elements. One approach to the
identification of channel domains that participate in gating is to
locate the sites of action of modifiers. Covalent reaction of Kv2.1
channels with the neutral, sulfhydryl-specific
methylmethanethiosulfonate (MMTS) caused a slowing of channel gating
with a predominant effect on the kinetics of activation. These effects
were also obtained after intracellular, but not extracellular,
application of a charged MMTS analog. Single channel analysis revealed
that MMTS acted primarily by prolonging the latency to first opening
without substantially affecting gating transitions after the channel
first opens and until it inactivates. To localize the channel
cysteine(s) with which MMTS reacts, we generated
NH2- and COOH-terminal deletion mutants and a construct in which all three cysteines in transmembrane regions were substituted. Only the
NH2-terminal deletion construct gave rise to currents that activated slowly and displayed
MMTS-insensitive kinetics. These results show that the
NH2-terminal tail of Kv2.1 participates in transitions leading to activation through interactions involving reduced cysteine(s) that can be modulated from the
cytoplasmic phase.
electrophysiology; site-directed mutagenesis; chemical
modification; cysteine
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INTRODUCTION |
VOLTAGE-DEPENDENT ION channels visit different
conformational states driven by changes in membrane potential.
Voltage-gated K+ channels reach
the open (active) state after depolarization and the closed (deactive)
state on repolarization. Additionally, they close (inactivate) in the
presence of sustained depolarizations by alternative mechanisms. The
transition rates between these states can be profoundly affected by
mutations in several channel domains. Extensive work has identified
regions involved in inactivation. Deletion of residues 6-46 of
Shaker
K+ channels abolishes the fast
inactivation observed in these channels (7), whereas mutations in the
sixth hydrophobic domain (S6) alter the rate of a separate slow
inactivation process (8). Amino acid substitutions in the pore-forming
(P) region of Kv2.1, a channel that inactivates slowly, speed or delay
inactivation (6; J. M. Pascual and A. M. Brown, unpublished
observations). On the other hand, evolutionarily conserved residues in
the S4 segment play a key role in activation gating and have been well characterized as sensing elements required for voltage-dependent activation (13, 17). A substantial part of the charge movement that
leads to channel opening can be accounted for by voltage-facilitated translocation of basic S4 residues relative to the protein core (15,
31). Analysis of the rates of reaction of hydrophilic methanethiosulfonate derivatives with substituted cysteines showed that S4 residues are subject to chemical modification from the extracellular or the cytoplasmic phase, depending on the state of the
channel gates (11, 30). These experiments support the notion that,
instead of being buried in the rest of the protein core, the voltage
sensor is exposed on both sides of the membrane and is available to
modulation from the cytoplasmic phase, as proposed by Perozo and
Bezanilla (20). Although other S4-interacting residues have been
identified in transmembrane areas of the channel (16), considerable
effort will be required to identify the rest of the structural
components that participate in voltage-dependent gating and to
elucidate how they regulate ion flow through the pore.
We set out to identify channel regions involved in
K+ channel activation outside S4.
Our search was motivated by the findings of Caputo et al. (4) using the
sulfhydryl-specific reagent p-hydroxymercuriphenylsulfonic acid
(PHMPS) on squid axon K+ channels.
PHMPS causes a pronounced slowing of activation gating, yet there are
no cysteines in the S4 region of cloned voltage-gated channels
(including the squid K+ channel
from optic lobe; D. Patton and F. Bezanilla, personal communication;
21), suggesting that PHMPS reacts with a target located elsewhere in
the channel itself or in a closely associated subunit. Our approach to
localizing the sulfhydryl-reactive domain(s) associated with gating
modification combined cysteine elimination by genetic substitution or
deletion with a functional assay for the loss of sensitivity to
sulfhydryl reagents. We have used the sulfhydryl modifier
methylmethanethiosulfonate
(CH3SO2SCH3,
MMTS) extracellularly applied on Kv2.1 expressed in
Xenopus oocytes. MMTS was chosen
because 1) it reacts specifically
with cysteinyl sulfhydryls, attaching a thiomethyl group via disulfide
bonding (24), 2) it is uncharged,
small, and lipid soluble, potentially having access to
membrane-embedded and cytoplasmic regions, even when applied
extracellularly, and 3) several
charged, hydrophilic MMTS analogs are available (1).
In whole cells, MMTS reacted with Kv2.1, producing an irreversible
slowing of activation kinetics, which was due to a prolongation of the
latency to first opening, as shown by single channel analysis. This
effect could be prevented by deleting the first 139 amino acids from
the NH2 terminus of the channel.
Additionally, MMTS decreased channel conductance by modification of a
separate site that could be regenerated by the reducing agent
dithiothreitol [HSCH2CH(OH)CH(OH)CH2SH,
DTT]. The results support the involvement of the
NH2 terminus of Kv2.1 in
voltage-dependent gating and its susceptibility to modulation from the
cytoplasm.
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MATERIALS AND METHODS |
Recombinant DNA.
Deletion of the 347 COOH-terminal amino acids (residues 506-853;
C) of Kv2.1 was achieved by the excision of a
BamH
I-Bgl I cassette followed by the
ligation of an adapter sequence coding for the amino acids RPPPEPER
followed by a stop codon. Incorporation of this epitopic sequence did
not affect unitary channel conductance or gating, as detailed by
Pascual et al. (19). Substitutions C232A, C393S, and C394S, located in
membrane-associated regions, were sequentially introduced on a
full-length Kv2.1 clone by oligonucleotide-mediated mutagenesis using a
phage-subcloned, single-stranded Kv2.1 DNA cassette. The
NH2-terminal deletion construct
(
N) was generated by digestion of Kv2.1 DNA with
Cla I, which released a fragment encoding the first 120 amino acids and most of the
5'-untranslated and polylinker regions. Self-ligation of the
remaining segment yielded a clone in which the first in-frame
methionine was at position 140, therefore lacking the first 139 amino
acids. Substitutions C163S and C164S were also performed by
oligonucleotide-directed mutagenesis. All constructs were verified by
restriction analysis and dideoxy sequencing extending over mutant and
ligation areas.
Capped runoff cRNA transcripts were prepared after plasmid
linearization with Not I, quantitated,
and stored in 100 mM KCl at
80°C, as described by Pascual et
al. (19). Stage V and VI oocytes were injected with 46 nl of
sufficiently concentrated cRNA to achieve 3-10 µA of whole cell
current at 40 mV. cRNA was injected (in ng/µl) as follows: 2 Kv2.1,
0.5
C, 120
N, 50 transmembrane core cysteineless, and 5 all other
mutants. Cells were maintained in culture for 1-5 days before
electrophysiological analysis.
Whole cell recording.
Cells were transferred to a 250-µl recording chamber and perfused at
5 ml/min with a solution including (in mM) 51 KOH, 60 NaOH, 120 methanesulfonic acid, 11.5 N-methyl-D-glucamine,
10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid (HEPES), and 2 CaCl2,
adjusted to pH 7.2. The high KOH concentration allowed the measurement
of large-amplitude inward tail currents on membrane repolarization. A 3 M KCl-agar bridge connected the recording chamber to an Ag-AgCl
reference electrode. The ethylammonium analog of MMTS (MTSEA,
CH3SO2SCH2CH2NH3+)
was a gift of Arthur Karlin. MMTS was purchased from Sigma Chemical (St. Louis, MO) and DTT from US Biochemical (Cleveland, OH). Because methanethiosulfonates hydrolyze rapidly at neutral pH and room temperature (A. Karlin, personal communication), MMTS and MTSEA were
dissolved in perfusion solution immediately before application to the
oocyte and used for a maximum of 5 min. Low-resistance (0.3-0.7
M
) agarose-cushion electrodes were manufactured from beveled glass
micropipette tips filled with a 0.5% agarose-3 M KCl gel and
backfilled with 3 M KCl. Currents were leak and linear capacitance
subtracted using an on-line P/4 routine. Similar results were obtained
in experiments performed without subtraction. Cells exposed to MMTS
were monitored for a decrease in membrane resistance, which was often
observed while the reagent was being perfused. Cells in which the leak
at
80 mV reached 10% of the voltage-gated ionic conductance at
40 mV or in which the leak persisted after withdrawal of MMTS were
discarded from the analysis. Macroscopic tail currents were elicited by
instant repolarizing voltage pulses to
80 mV after prolonged
depolarizing prepulses of sufficient duration (0.5-1.5 s) to
activate most channels. Tail current amplitudes were measured by
extrapolation of monoexponential fits back to the time of the voltage
jump. Steady-state activation gating was analyzed by fitting
current-voltage (I-V) relationships
obtained from tail current measurements in whole oocytes. Tail current amplitudes were fitted to the Boltzmann equation:
I/Imaximum = {1 + exp[(V1/2
Vm)/k]}
1,
where I represents normalized tail
current amplitude,
Vm is prepulse
potential, V1/2
is activation midvoltage, and k is a
slope parameter expressed in F/RT
units. Experiments were carried out at 21-23°C. Values are
means ± SE. A two-tailed t-test was used to assess the significance of the difference between means.
Giant patch recording.
Patch pipettes of ~100 µm diameter were polished to ~0.5 M
of
resistance when filled with (in mM) 120 NaCl, 2.5 KCl, 2 CaCl2, and 10 HEPES, pH 7.2. Bath
solution containing (in mM) 100 KCl, 10 EGTA, and 10 HEPES, pH 7.2, was
used to zero the membrane resting potential. Inside-out patches were
excised from mechanically devitellinized oocytes, and their
intracellular aspect was exposed to bath solution. An Axopatch-1D
amplifier (Axon Instruments, Foster City, CA) was used. Currents were
low-pass filtered at 0.5-1 kHz (
3 dB, 4-pole Bessel filter)
and digitized at a sampling rate of 2 kHz. Capacitance was compensated.
Leak currents and series resistance were negligible and were left
uncompensated. Typically, initial giant patch peak currents measured
from oocytes expressing Kv2.1 reached 1-4 nA and decayed
monoexponentially after repetitive depolarization to a steady level
with a time constant of ~3 min at a holding potential of
80
mV, as described by Pascual et al. (19). Exceptionally, a few patches
did not exhibit rundown. The effects of internal MTSEA application to
giant patches did not depend on rundown velocity or magnitude.
Single channel recording.
Cell-attached single channel activity was recorded using the equipment
and solutions described for giant patch experiments, except pipettes
were fire polished to a resistance of 5-10 M
. Channels were
activated with test pulses of 0-40 mV from a holding potential of
60 or
80 mV. Currents were low-pass filtered at 1 kHz
before digitization at 4 kHz. Records were corrected for linear
capacitive and leak currents by subtracting the smoothed average of
records lacking channel activity (10). Single channel currents were
idealized using TRANSIT (27). This algorithm uses a
dI/dt
threshold to identify transitions between open and closed levels during
a first data pass. In a second pass, idealized openings were
constructed using a half-amplitude threshold criterion. Amplitudes and
open and closed interval durations of idealized data were collected
into distribution histograms. Amplitude histograms were fitted with
Gaussian functions using a maximum likelihood estimate. Slope
conductances were determined from least-squares fits of current
amplitude-voltage data in a test potential range of 0-60 mV. Mean
open times
(topen) were
calculated by fitting open time histograms to a single-exponential
decay function using a maximum likelihood estimate. Open times <0.4
ms were excluded from fitting. The distribution histograms of closed
interval durations were fitted with two-exponential decay functions
from which mean closed times within a burst
(tclosed) or
between bursts
(tinterburst) were determined. Bursts were defined by establishing a critical closed
interval of 1 ms, and their mean durations
(tburst) were obtained by fitting the duration histograms to a single exponential. The time constant of the first latency (waiting time from the start of
the test pulse to first opening) was deduced from monoexponential fits
to cumulative distribution histograms.
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RESULTS |
Currents arising from the expression of Kv2.1 in oocytes activate over
several milliseconds following a sigmoidal time course, a feature of
delayed rectifier channels, and display a slow inactivation process
appreciable at depolarized voltages (>20 mV; Fig.
1A). The channels deactivate on return to hyperpolarized potentials, generating inward tail currents, which are noticeable because of the
high extracellular K+
concentration used in this experiment (see MATERIALS
AND METHODS). Activation of MMTS-treated channels
required larger depolarizations and followed a slower time course than
Kv2.1 control currents (Fig. 1B). A
parallel phenomenon was a prolongation of the time course of channel
deactivation, as indicated by the slowing of tail current decays. These
effects developed slowly, over several minutes of MMTS application, and
could not be reversed by washout, as expected if MMTS reacted
covalently with sulfhydryl groups associated with the channel.

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Fig. 1.
Modification of Kv2.1 currents by sulfhydryl reagents.
A: whole oocyte control currents
elicited by a series of depolarizing potentials in 10-mV steps from
holding potential of 80 mV. B:
application of 4 mM methylmethanethiosulfonate (MMTS) over 5-min
prolonged time course of activation and delayed deactivation tail
currents. C: subsequent superfusion of
5 mM dithiothreitol (DTT) over 5 min caused slight increase in current
amplitudes but had no effect on kinetics. Reactions were monitored to
completion.
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To test the accessibility of the site(s) modified by MMTS, we attempted
to reverse the reaction by reduction with DTT. In control experiments,
high (50 mM) concentrations of DTT had no effect on Kv2.1 currents. We
reasoned that if MMTS reacted with a single, readily accessible site,
DTT would regenerate the cysteinyl sulfhydryl, gradually restoring
currents to their premodification appearance. However, application of 5 mM DTT to MMTS-treated cells increased current amplitudes only
slightly, with no effect on activation or deactivation kinetics (Fig.
1C). This result suggests that
access of the slightly larger DTT to the MMTS-modified site that causes
alteration of channel kinetics is restricted and/or that
penetration of DTT through the bilayer to accumulate in the cytoplasmic
phase proceeds slowly. Scaled MMTS-modified channel currents
superimposed on post-DTT current traces (not shown), indicating that
gating follows an identical time course under both conditions. This
result is consistent with a dual action of MMTS:
1) a predominant effect on channel
kinetics, irreversible under our experimental conditions, and
2) a small decrease in conductance
that arises from modification of an additional site that can be
subsequently reduced by DTT.
Effect of MMTS on activation kinetics.
To describe the changes in channel kinetics after MMTS in quantitative
terms, we measured the time course of current activation in whole
oocytes. As noted above, activation of Kv2.1 currents after a 40-mV
depolarizing test pulse proceeds following a sigmoidal time course
(Fig. 1A). This phenomenon is
compatible with the contention that the channels must traverse several
closed states before opening. We fitted the 5-95% amplitude
interval of these currents with the sum of two exponentials of similar
amplitudes and time constants (
) of 11 ms
(
fast) and 24 ms
(
slow) (Table 1). This procedure eliminated the foot of
the activation process from the fit, which arises from channels that
open early during depolarization. On reaction with MMTS (Fig.
1B), the slower exponential component of activation was substantially affected, becoming prolonged by fourfold
(
slow MMTS = 95 ms), whereas the fast component was only marginally slowed
(
fast MMTS= 16 ms; Table
1). This effect persisted in oocytes expressing Kv2.1 and treated with
DTT after reaction with MMTS (not shown), as if the site that caused
conductance reduction after
S-methylation could be modified and
regenerated independently of the site(s) that mediated the effects of
MMTS on channel kinetics. Another action of MMTS was a prolongation of
the time course of deactivation, as measured by fitting tail currents
with single-exponential functions. As shown in Fig. 1, after MMTS
treatment, Kv2.1 channels deactivate more slowly (deactivation
= 19 ms) than in control conditions (deactivation
= 7 ms), indicating
that open channels remain open longer before they close. This effect
also persisted after DTT application (deactivation
= 18 ms).
We also determined the voltage dependence of channel activation
(I-V relationship) in the unmodified
Kv2.1 (Fig.
2A). The V1/2 for
steady-state activation of Kv2.1 currents, obtained by fitting
I-V data to a Boltzmann equation (see
MATERIALS AND METHODS), is
1
mV, with a k of 11 F/RT (Table 1). In contrast, activation of MMTS-treated Kv2.1 channels proceeded too slowly to reach
steady state at the end of test potentials of sufficient duration to
fully activate Kv2.1 in control conditions (Fig. 1). Prolongation of
test pulses impaired clamp performance in these cells and prevented
accurate steady-state measurements. Therefore, we could not measure the
shift of the activation midpoint of these channels.

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Fig. 2.
Effect of internal application of ethylammonium analog of MMTS (MTSEA)
on Kv2.1. Cytoplasmic aspect of an inside-out giant patch was exposed
to 2 mM MTSEA for 2 min. Currents were elicited by test pulses to 40 mV
from holding potential of 80 mV. A patch with exceptionally
stable currents is shown. Similar results were obtained in 5 other
patches presenting various degrees of current rundown.
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Sidedness of MMTS reaction.
We began to search for the structural element(s) responsible for the
kinetic changes induced by MMTS by investigating the pathway from which
MMTS gained access to its target in the channel. The neutral character
of MMTS makes it potentially membrane permeant. Therefore, reaction
with Kv2.1 could take place 1) from
the extracellular solution, 2) from
the membrane phase with MMTS molecules that partition in the lipid
bilayer, or 3) after diffusion
through the membrane and intracellular accumulation before the
molecules gained access to cytoplasm-exposed areas of the channel. From a simple perspective, if the reaction occurred from the
extracellular phase with a solvent-exposed site, we would expect
it to proceed at a relatively fast rate, whereas subsequent partition
in the bilayer and/or penetration through protein-protein
interfaces would impose a diffusional barrier that could be expected to
slow the observed rate of reaction. Similarly, the reducing environment provided by intracellular compounds such as glutathione would also
retard access of MMTS to Kv2.1 if the reaction took place in the
cytoplasmic phase.
A transient increase in oocyte resting conductance during MMTS
perfusion prevented accurate determination of channel modification rates. Nevertheless, high (4 mM) MMTS concentrations applied for at
least 2 min were required to drive the reaction to completion. This
result stands in contrast with that obtained for the reaction of MMTS
with 2-mercaptoethanol, which appears to proceed two to three orders of
magnitude faster, as determined by Stauffer and Karlin (25). Therefore,
MMTS modified Kv2.1 at a much slower rate, as if access to the reactive
cysteine(s) was restricted by a diffusional barrier or as if MMTS was
being quenched by other reactive groups in the oocyte interior.
However, an alternative possibility was that the reaction rate-limiting
step was not access to the site but disulfide formation. This would be
the case if, for example, the target site(s) was extracellularly
accessible but had a poor intrinsic reactivity because of infrequent
ionization of the cysteinyl sulfhydryl. Support for either of the
former hypotheses vs. the latter was provided by experiments using
MTSEA. MTSEA is hydrophilic and mostly charged at pH 7.2, being less permeant through the lipid bilayer than MMTS. Therefore, fast reaction
with MTSEA is indicative of solvent exposure (1). Whereas extracellular
application of MTSEA (up to 5 mM) and other charged MMTS
analogs had no effect on Kv2.1 currents (19), exposure of the
intracellular side of giant patches containing numerous channels to 2 mM MTSEA resulted in an irreversible slowing of activation kinetics
that developed nearly instantaneously and resembled the effects of MMTS
applied to saturation by superfusion (Fig. 2). Therefore, the site
responsible for modulation of Kv2.1 kinetics appeared to be accessible
to MTSEA from the cytoplasmic but not from the extracellular phase.
Locating a target domain for MMTS.
We continued the search for the channel region mediating the effects of
MMTS by assigning the 15 cysteines present in the Kv2.1 clone to 3 primary structural domains: 4 in a COOH-terminal domain, 3 in a
transmembrane core region (including membrane-related segments
S1-S6), and the remaining 8 in an
NH2-terminal domain. To identify
on which of the channel domains MMTS reacted to modify gating, we
engineered three cysteine-deficient constructs (Fig. 3): 1) a
COOH-terminal deletion that eliminates the last 347 residues (amino
acids 506-853), which included all COOH-terminal cysteines (
C),
2) a transmembrane core cysteineless
construct combining substitutions C232A (located in S2) and C393S and
C394S (both in S6), and 3) an
NH2-terminal deletion by excision
of the first 139 amino acids (
N). This deletion mutant lacked the
first six cysteines. Several other single and multiple cysteine
substitutions were also studied. A list of these mutants, together with
some of their kinetic properties, is given in Table 1.

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Fig. 3.
Effects of MMTS on cysteine-deficient Kv2.1 mutants.
Top: division of Kv2.1 into primary
structure domains. S1-S6, putative membrane-spanning segments; P,
pore-forming region. Location of cysteines [at positions 25, 59, 69, 107, 128, 129, 163, 164 (NH2
terminus); 232, 393, 394 (core region); 590, 705, 808, 826 (COOH
terminus)] is indicated by C. A, alanine; S, serine.
A and
B: COOH-terminal deletion ( C) and
transmembrane core cysteineless channels are susceptible to
modification by MMTS. Post-MMTS current trace crosses over
unmodified current record because this construct lacks all
transmembrane cysteines, which are responsible for a reduction in
conductance observed in Kv2.1. C: MMTS reduced current
amplitude on N channels but did not affect current kinetics.
Inset: superimposition of current
traces from N (control) and MMTS-treated channels (scaled after
multiplication by 1.165). Cells were depolarized to 40 mV from holding
potential of 80 mV. Calibration bars: 50 ms, 0.5 µA.
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All mutant constructs produced robust (several µA), voltage-gated
K+ currents in oocytes suitable
for detecting changes in kinetics after MMTS exposure. Activation of
COOH-terminal deletion and transmembrane core cysteineless channels
resembled Kv2.1, except for a negative shift
(V1/2 =
20
mV) in the voltage dependence of activation of cysteineless core
channels, which could be largely accounted for by the substitution
C232A carried by that construct (Table 1). Nevertheless, as illustrated
in Fig. 3, A and
B, activation induced by strong
depolarizing pulses was prolonged for both of these mutants after
reaction with MMTS, indicating that the target cysteine(s) associated
with the gating machinery remained unaltered by these mutations and
therefore must be located in the
NH2 terminus or, perhaps, in an
intimately associated subunit that interacts with the
NH2 terminus of the channel.
Indeed, deletion of the first 139 residues generated channels that
activated over a prolonged time interval, displaying kinetics that were
unaffected by MMTS application (Fig.
3C, Table 1). The time course of
activation of unmodified and MMTS-treated
N currents overlapped that
measured for MMTS-modified Kv2.1 channels (Table 1). Analysis of
N
steady-state activation gating also supported the conclusion that the
effects of MMTS on Kv2.1 are mediated by reaction with the
NH2 terminus. The
V1/2 of
N
currents was 53 mV, with k of 13 F/RT (Table 1). After reaction with
MMTS, both parameters remained unchanged (Table 1, Fig.
4), despite a reduction in channel
conductance, as described below. Replacement of residues C163 and C164
(located before S1 and left in place in the
N deletion construct)
with serine had no effect on the time course of activation, affecting only marginally its voltage dependence (Table 1), and was not analyzed
further. Nevertheless, MMTS still reacted with
N channels, causing a
slight (15%) decrease in current amplitude, an observation that is
consistent with the hypothesis stated above of an additional site that
is responsible for the reduction in conductance noticed after
modification and appears to remain intact in the deletion mutant.
Compensating for current reduction by multiplying amplitudes by a
scaling factor (ranging from 1.12 to 1.19 in 10 cells tested) allowed
accurate superimposition of
N current onset traces obtained before
and after MMTS application (Fig. 3,
inset), supporting the insensitivity
of
N channel gating to MMTS. Therefore, both MMTS targets appear to
modulate different aspects of channel function (kinetics and
conductance) from separate loci.

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Fig. 4.
Steady-state current-voltage relationships in N channels.
A: control conditions before reaction
with MMTS. Tail currents were normalized to maximum current (set to 1),
averaged, fitted with a Boltzmann equation, and plotted against
prepulse potential (solid curve; see MATERIALS AND
METHODS). B: N
current-voltage data obtained after modification by MMTS. Conductances
were normalized to 1 as in A.
Solid-line fit has been replotted from
A to illustrate similarity in
steady-state activation gating in N channels before and after MMTS
reaction. Each set of symbols represents an independent experiment.
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Conduction through mutant and modified channels.
Measurement of instantaneous I-V
relationships allowed the assessment of ion conduction in open Kv2.1
and
N channels before and after MMTS treatment. As shown in Fig.
5, Kv2.1 currents display outward
rectification in asymmetric solutions (51 and 60 mM extracellular KOH
and NaOH, respectively) (see MATERIALS AND
METHODS; 19). Kv2.1 and
N channel currents showed a
similar degree of rectification, a phenomenon that is consistent with
the integrity of the permeation pathway in
N channels. Figure 5 also
illustrates the effects of MMTS modification on ion conduction through
N channels. After MMTS reaction, outward currents were slightly more
reduced than inward currents, without shifting reversal potential,
resulting in a smoothing of the rectifying profile of
N channels.

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Fig. 5.
Instantaneous current-voltage relationships in Kv2.1, N, and
modified channels. A and
C: whole cell current recordings
obtained from oocytes expressing Kv2.1 and N channels, respectively.
B and
D: averaged current-voltage plots from
5-12 oocytes. D also shows effect
of modification of N currents. Reaction with MMTS reduced inward and
outward conductance, causing a slight reduction of rectification in
N channels. Oocytes were depolarized to test potentials increasing
in 10-mV intervals after a constant activating prepulse to 20 (Kv2.1)
or 40 mV ( N) from holding potential of 80 mV. Unmodified
current amplitudes were normalized to amplitudes measured at 50 mV (set
to 1).
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Figure 6 and Table
2 illustrate the effects of MMTS
modification on currents through single channels and their
correspondence with the whole cell measurements described above. After
MMTS reaction, the unitary slope conductance of Kv2.1 measured in a
range of positive potentials significantly decreased from 10 to 8.2 pS (P < 0.05). This result correlated
with and accounted for the current reduction (~15%) obtained from
the outward limb of macroscopic I-V
relationship measurements. On the other hand, the conductance of single
N channels (9.2 pS) was not significantly different from Kv2.1, in
support of the notion that the
NH2-terminal deletion does not
interfere with K+ permeation.

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Fig. 6.
Single channel analysis of Kv2.1, modified, and N channels.
A-C: cell-attached single channel
currents recorded at 40-mV test potentials from holding potential of
60 mV. Dotted line, baseline. Calibration bars: 1 pA, 100 ms.
D: cumulative mean first latency
distributions obtained by plotting normalized cumulative channel open
probability at 40 mV against channel waiting time and fitted with a
single exponential. E: test voltage
dependence of mean first latency. Data were obtained from 2-10
patches.
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Mechanism of gating modification by MMTS.
Analysis of single channel kinetics yielded information about changes
in Kv2.1 gating phenotype after reaction with MMTS (Fig. 6, Table 2).
Channel kinetics were studied in the presence of 400-ms depolarizing
pulses to minimize transit to the inactivated state, which occurs at a
rate of 0.3 s
1, as measured
in whole cell recordings. Five parameters were used to characterize
gating of individual Kv2.1 channels:
1) latency to first opening after a
depolarizing step, 2) burst duration (defined as a family of related openings interrupted only by brief closings;
tburst),
3) interburst interval
(tinterburst),
4) open time
(topen), and
5) closed time (including all time
intervals that the channels spend closed except
parameters 1 and
3,
tclosed). Average values for these parameters are given in Table 2 for Kv2.1,
MMTS-treated Kv2.1, and
N channels, and representative recordings
are shown in Fig. 6.
Remarkably, only the first latency appeared to be significantly
prolonged (~4-fold) in MMTS-exposed and
N channels compared with
Kv2.1. None of the other parameters measured diverged significantly from Kv2.1 values (Table 2). These results indicate that the conformational changes underlying the transitions responsible for the
behavior of the channel after the first opening are unaffected by MMTS
or by the deletion of the
NH2-terminal domain. Figure 6D plots the cumulative probability of
first opening vs. time of Kv2.1, MMTS-treated, and
N channels after
a voltage step to 40 mV. As also observed in
Shaker channels (2, 9), the sigmoidal appearance of the three plots resembles the time course of macroscopic activation and further indicates the existence of several closed states
that can be visited before opening. Additional support for the
similarity of functional effects between MMTS modification and
NH2-terminal deletion on the
kinetics of activation is provided by their similar first latency
distributions, which stands in contrast to the steeper distribution
obtained for Kv2.1 (Fig. 6D).
As exemplified by noninactivating
Shaker channels (2, 9), the latency of
activation of delayed rectifiers is voltage dependent. At depolarized
test potentials, the latency to first opening decreases as the channels
travel faster through the activation pathway. Figure
6E illustrates the voltage dependence
of the latency to first opening of Kv2.1 channels. As expected,
depolarizing test pulses of increasing magnitude shortened the delay of
first Kv2.1 openings. A similar voltage dependence was appreciated in the latencies of MMTS-treated Kv2.1 and
N channels, despite their prolonged values, suggesting that, over the potential range studied, the process by which MMTS and the deletion of the
NH2 terminus affected activation
has little intrinsic voltage dependence.
An additional effect of MMTS was a slowing of deactivation, which was
observed in tail current measurements (Fig. 1). This phenomenon
suggests that reaction with MMTS also increases the stability of the
open state at negative potentials, perhaps by slowing transitions
leading away from the activated state.
Effect on inactivation gating.
Inactivation of Kv2.1 currents proceeds slowly over several seconds and
can be well approximated with a single-exponential function with a time
constant of 3.3 s. MMTS decreased the rate of inactivation of Kv2.1
channels, as revealed by long depolarizing pulses (Fig.
7). Inactivation time constants increased
for Kv2.1 and transmembrane core cysteineless currents after
application of MMTS. Similar results were obtained in oocytes
expressing
C channels (not shown). In contrast,
N currents
displayed little inactivation under prolonged depolarizations (up to 20 s) and were only scaled down by MMTS (Fig. 7). Therefore, these
experiments do not indicate whether the effects on Kv2.1 inactivation
were also mediated through modification of the
NH2 terminus or arose from
reaction with cysteine(s) located in another domain. Nevertheless, the
observation that the
N deletion abolished inactivation suggests a
central role of the NH2 terminus
in the slow inactivation of Kv2.1, as previously noted (28).

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|
Fig. 7.
Inactivation of Kv2.1 and mutant channels after MMTS treatment.
A and
B: MMTS slowed inactivation in
constructs carrying intact NH2
terminus. Similar results were obtained in C channels.
C: deletion of
NH2-terminus generated channels
with inactivation kinetics unaffected by MMTS. These channels showed no
noticeable inactivation, even during test pulses lasting 60 s. A small
reduction in current amplitudes after reaction with MMTS is apparent.
Calibration bars: 2 s, 2 µA.
|
|
 |
DISCUSSION |
Effects of Kv2.1 sulfhydryl modification.
Reaction of Kv2.1 with cysteine-specific MMTS caused a pronounced and
irreversible slowing of channel kinetics. We centered our studies on
the modulation of activation by MMTS, because
1) all Kv2.1 cysteines are located
outside S4, a central component of the voltage sensor, indicating that
other channel regions not yet identified must be involved in the
effects of MMTS and may, therefore, participate in gating, and
2) as summarized by Caputo et al.
(4), most studies on chemical modification of native excitable
membranes using sulfhydryl reagents report (somewhat discrepant)
results on K+ channel
inactivation, with less emphasis on activation. The findings presented
suggest that MMTS modulates activation of Kv2.1 by reacting with one or
more cysteines located at the NH2
terminus. Sequential cysteine-elimination mutagenesis results together
with functional assay show that the presence of the hydrophilic
NH2-terminal domain, which has
been shown to be intracellular (29), is required for MMTS
susceptibility. An alternative possibility is that the reactive cysteine(s) is located in an accessory or modulatory subunit that is
present in the oocyte and associates with Kv2.1. However, there is no
evidence that Kv2.1 interacts with other subunits when expressed in
oocytes. Therefore, we favor the proposal that the effects of MMTS are
due to direct reaction with the
NH2 terminus of the channel. On
the basis of the slowness of the reaction when MMTS is extracellularly
applied and the finding that a charged MMTS homolog (MTSEA) readily
reacts when applied to the intracellular but not the extracellular side
of the membrane, we infer that the
NH2-terminal target is exposed to
and preferentially accessible from the cytoplasmic phase.
VanDongen et al. (28) reported that extensive deletions at the
NH2 terminus of Kv2.1 were
associated with marked changes in whole cell current kinetics that were
concomitant with a drastic decrease in current expression levels,
raising the concern that the structural integrity of the channel had
been globally affected. The single channel analysis results reported
here, however, disprove that possibility.
NH2-terminal deletion and chemical
modification by MMTS cause specific and similar changes in a set of
channel properties but preserve all others, indicating that
1) the overall structure of the
protein is most likely unchanged in both cases and
2) the effects of deleting the
NH2-terminal domain resemble those
obtained by modification of reduced cysteine(s) located in it. Because
single channel conductance and open probability after first opening are
largely preserved, we conclude that the reduced whole cell current
levels observed with the
N construct arise from a reduction in the
number of channels that populate the plasma membrane.
The mechanism responsible for the delay of activation observed in Kv2.1
channels after reaction with MMTS and the slow activation of
N
channels appears to be an increase in the latency to first opening,
which was invariant over the potential range studied. Open and closed
times after first opening and before inactivation were virtually
unaffected by MMTS. Therefore, in the presence of depolarizing
potentials, alterations at the
NH2-terminal domain influence only
transitions in the activation pathway before first opening (2, 8) but
have little influence after opening and until the channel sojourns in
the inactivated state. The fact that in MMTS-treated channels the
latency to first opening was prolonged regardless of voltage may
suggest that MMTS modulates a voltage-insensitive channel transition,
but it is also possible that MMTS slows the rates of interconversion
between two or more states in voltage-dependent segments of the
activation pathway of the channel. In either case, the modification
does not seem to affect the stability of the fully activated channel
while depolarization is maintained. On the other hand, the observed
slowing of tail currents under repolarizing potentials induced by MMTS
reaction indicates that at negative potentials the open state of the
channel has been stabilized. Further channel kinetic analyses are
required to characterize the steps in the activation pathway of Kv2.1
and the manner in which MMTS affects their rates of interconversion by
reacting with the NH2 terminus.
A separate effect of MMTS was a small decrease in current amplitude,
which was verified by single channel conductance measurements and could
be restored by reaction with the reducing agent DTT. The conductance of
Kv2.1 and
N channels was similar in control conditions (10.0 vs. 9.2 pS). MMTS reduced the single channel conductance of Kv2.1 by 15%,
whereas a similar reduction was observed after analysis of MMTS-treated
N whole cell currents, despite the lack of an effect on channel
kinetics. Additional support for the similarities in structure and
function of both channel pores is provided by their analogous
instantaneous I-V profiles. Taken
together, these results indicate that a minimum of two cysteines are
involved in MMTS action on Kv2.1: one associated with the NH2 terminus and accessible from
the cytoplasm and another outside the
NH2 terminus, which can react with
DTT after modification by MMTS and possibly lines the conduction
pathway. A likely candidate for the latter residue is C393, which is
located in S6 and has been proposed to lie near the pore (33).
Evidence for a dual effect of oxidizing agents on delayed rectifier
currents was also provided by the experiments of Caputo et al. (4) on
native squid axon channels using PHMPS. PHMPS causes a slowing of
channel kinetics and a reduction in single channel conductance that are
remarkably similar to our findings with cloned Kv2.1. In addition, a
decrease in open probability accompanies these effects. Although the
latter phenomenon was not observed in Kv2.1, we propose that the
NH2-terminal domain of squid
K+ channels may be responsible for
the modulatory effects of PHMPS on channel kinetics and that it plays a
similar role in activation, and we infer that participation of the
NH2-terminal domain in activation
gating is a generic feature of Kv2-related channels. In contrast,
cysteine replacement (3), MMTS exposure (14), or deletions (7, 26) at
the NH2 terminus of
Shaker channels do not affect
activation gating, suggesting a divergent functional specialization.
Mutational evidence argues against the involvement of disulfide bonding
between NH2-terminal residues and
the rest of the channel protein as the mechanism underlying gating
modulation. According to that view, hypothesized by Ruppersberg et al.
(22), oxidation of cysteine residues at the
NH2-terminal segment involved in
fast inactivation of A-type K+
channels would uncouple the inactivating domain from its receptor by
preventing the formation of cystine. Although a significant fraction of
the cysteines in solved structures is found bridging to remote sites in
the primary amino acid sequence (5), in our experiments with Kv2.1,
point or combined elimination of all cysteines in regions downstream of
the NH2 terminus did not have significant effects on channel activation or on the ability of MMTS to
affect gating. These results suggest that disulfide bonding to residues
located elsewhere in the channel protein is not required for the
participation of the NH2 terminus
in gating. The observation that MMTS and MTSEA produce similar effects,
despite their differently charged attaching groups (0 and +1,
respectively), raises the possibility that the interference of modified
cysteines with the gating process is caused by steric hindrance in a
region subject to a conformational rearrangement during gating.
Contribution of the NH2-terminal domain
to K+ channel
function.
The two hallmark aspects of voltage-gated channel function are
selective ion conduction and control of gating by membrane potential.
The developments brought about by mutational and functional analysis in
the past 10 years have led to the notion that both properties are
specified in evolutionarily conserved protein motifs. Whereas ion
selectivity appears to arise from interactions with amino acids in the
P region, voltage-dependent gating requires the presence of the
amphipatic domain S4. Conversely, the modulation of these properties
and processing of channel subunits by the cell reside in less conserved
regions, allowing for a wide phenotypic diversity.
The NH2-terminal domain of
voltage-gated K+ channels is
variable in length and amino acid composition. Nevertheless, key roles are specified in it. The compatibility of
NH2-terminal (together with other)
regions allows subunit association to form functional tetrameric
channels (23, 29). Heteromeric assembly of subunits with compatible
recognition domains is a mechanism used by cells to diversify
K+ currents. Fast channel closing
by N-type inactivation is provided by specialization of the first
residues of numerous K+ channels
into a tethered pore-blocking particle (7). The presence of
intracellularly exposed cysteine residues associated with the blocking
particle potentially allows for modulation by the redox state of the
cytoplasmic phase, a phenomenon that may have physiological relevance
(22). Additionally, the NH2
terminus of Shaker-related channels
provides a recognition region for modulatory (
) subunits that affect
the kinetics of channel inactivation (32).
Our findings illustrate a different aspect of the regulation of
K+ channel activity by the
NH2-terminal domain: modulation of
activation. Recent results that voltage-sensing residues in S4 are
accessible from the cytoplasmic phase (11, 31) and that charge transfer by phosphorylation offsets voltage-dependent parameters of channel gating (20) indicate that the membrane electric field sensor is
susceptible to modulation from the cytoplasmic compartment. Our
conclusion that the participation of the
NH2 terminus of Kv2.1 in gating
can be altered by chemical modification from the cytoplasmic phase is
best explained in the context of a gating apparatus involving several
domains in addition to S4 and subject to intracellular modulation.
 |
ACKNOWLEDGEMENTS |
We are indebted to Arthur Karlin for generosity and suggestions on
the manuscript. We thank Myles Akabas, Francisco Bezanilla, Arthur
Karlin, David Patton, and Antonius VanDongen for discussion of the
results and Wei-Qiang Dong for oocyte culture and injection.
 |
FOOTNOTES |
J. M. Pascual was supported in part by National Institutes of Health
Neurological Sciences Academic Development Award NS-01698. C.-C. Shieh
was funded in part by a grant-in-aid from the American Heart
Association, Northeast Ohio Affiliate. This work was also supported by
National Institute of Neurological Disorders and Stroke Grants NS-29473
to G. E. Kirsch and NS-23887 to A. M. Brown.
A preliminary account of these results has been reported in abstract
form (18).
Address for reprint requests: J. M. Pascual, Department of Neurology,
Box 164, Neurological Institute of New York, 710 W. 168th St., New
York, NY 10032.
Received 29 April 1997; accepted in final form 25 July 1997.
 |
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