From the Committee on Neurobiology, The University of Chicago, Chicago, Illinois 60637
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ABSTRACT |
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Site-3 toxins isolated from several species of scorpion and sea anemone bind to voltage-gated Na channels and prolong the time course of INa by interfering with inactivation with little or no effect on activation, effects that have similarities to those produced by genetic diseases in skeletal muscle (myotonias and periodic paralysis) and heart (long QT syndrome). Some published reports have also reported the presence of a noninactivating persistent current in site-3 toxin-treated cells. We have used the high affinity site-3 toxin Anthopleurin B to study the kinetics of this current and to evaluate kinetic differences between cardiac (in RT4-B8 cells) and neuronal (in N1E-115 cells) Na channels. By reverse transcription-PCR from N1E-115 cell RNA multiple Na channel transcripts were detected; most often isolated were sequences homologous to rBrII, although at low frequency sequences homologous to rPN1 and rBrIII were also detected. Toxin treatment induced a voltage-dependent plateau current in both isoforms for which the relative amplitude (plateau current/peak current) approached a constant value with depolarization, although the magnitude was much greater for neuronal (17%) than cardiac (5%) INa. Cell-attached patch recordings revealed distinct quantitative differences in open times and burst durations between isoforms, but for both isoforms the plateau current comprised discrete bursts separated by quiescent periods, consistent with toxin induction of an increase in the rate of recovery from inactivation rather than a modal failure of inactivation. In accord with this hypothesis, toxin increased the rate of whole-cell recovery at all tested voltages. Moreover, experimental data support a model whereby recovery at negative voltages is augmented through closed states rather than through the open state. We conclude that site-3 toxins produce qualitatively similar effects in cardiac and neuronal channels and discuss implications for channel kinetics.
Key words: patch clamp; voltage clamp; sea anemone toxin; cnidarian venoms ![]() |
INTRODUCTION |
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Insights into the molecular basis of Na channel inactivation have been achieved through a combination of
targeted structure/function studies (Fozzard and Hanck,
1996) and investigation of natural mutagenesis in a variety of genetic diseases that target Na channel inactivation in skeletal muscle (e.g., myotonias and periodic paralysis; Cannon, 1996
) and the heart (long QT syndrome; Ackerman, 1998
). Another important tool for
increasing our understanding of inactivation in Na
channels is the family of site-3 toxins (Catterall, 1980
),
which comprise a structurally diverse group of peptide
toxins isolated from several species of scorpion (Catterall, 1977
; Wang and Strichartz, 1983
) and sea anemone
(Narahashi et al., 1969
; Bergman et al., 1976
; Romey
et al., 1976
). These toxins bind to the extracellular face
of voltage-gated Na channels and prolong the time
course of sodium current (INa) by interfering with
channel inactivation (Narahashi et al., 1969
; Bergman
et al., 1976
; Romey et al., 1976
). Single-channel studies
(Schreibmayer et al., 1987
; Nagy, 1988
; Kirsch et al.,
1989
; El-Sherif et al., 1992
) and whole-cell studies
(Hanck and Sheets, 1995
) suggest that the prolongation of current arises from a selective inhibition of
channel inactivation from the open state, O
I, with
little effect on the activation process or on inactivation
from closed states (C
I), and a gating current study
in cardiac channels demonstrated that toxin binding
eliminates the charge movement associated with O
I
transitions (Sheets and Hanck, 1995
).
It is unlikely that the binding of toxin alters only a
single rate constant if this rate constant were located in
a kinetic loop, since such a modification would violate
microscopic reversibility. In addition, ionic current
studies have reported that toxin induces a low-amplitude noninactivating current (Bergman et al., 1976; Siemen and Vogel, 1983
; Gonoi et al., 1984
; Wasserstrom
et al., 1993
), as reflected by the failure of steady state
availability relations to reach zero, even at positive potentials. More interestingly, several studies using scorpion toxins have noted a noninactivating current component that increased in magnitude with increasing depolarization (Koppenhofer and Schmidt, 1968
;
Mozhayeva et al., 1980
; Wang and Strichartz, 1985
).
That is, steady state availability reached a nadir at an intermediate voltage and increased with further depolarization. Conversely, other investigations have reported
steady state availability relations that suggest that voltage-dependent inactivation is complete (Hanck and
Sheets, 1995
).
The lack of agreement between studies about the
presence and the magnitude of a plateau current could
be due to the differences among channel isoforms
studied. Neuronal Na channel isoforms have an O I
inactivation rate constant that is largely voltage independent (Aldrich et al., 1983
; Vandenberg and Bezanilla, 1991
). On the other hand, open-state inactivation
in cardiac channels is voltage dependent (Berman et al.,
1989
; Yue et al., 1989
; Scanley et al., 1990
) and is accompanied by a gating charge movement that is immobilized by site-3 toxins (Sheets and Hanck, 1995
). Differences in the observed effects of toxins may, therefore,
be secondary to kinetic differences in the channels they
modify. Alternately, the observed biphasic availability
relations might be peculiar to scorpion toxins. Finally,
differences might reflect specific discrepancies in the
voltage protocols and analysis methods used.
In these experiments, we studied the kinetic mechanisms underlying this plateau current. In particular, we
tested whether the noninactivating current arose from
a change in single-channel conductance, a persistent
rate of recovery from inactivation, or a toxin-induced
modal shift in channel gating. To address the issue of
channel isoform specificity, we recorded from neuronal channels expressed in the neuroblastoma cell
line N1E-115 and from cardiac channels expressed in
the cell line RT4-B8 (Zeng et al., 1996).
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METHODS |
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Cells and Cell Culture
Cells were grown in Dulbecco's Modified Eagle's Medium
(GIBCO BRL) supplemented with 9% fetal bovine serum and
were maintained at 37°C in a humid atmosphere containing 5%
CO2. Before recording, cells were dissociated from their substrate by brief (<2 min) digestion in a solution containing 0.25%
trypsin with 1 mM EDTA. Cardiac INa was recorded from the rat
peripheral neurotumor cell line RT4-B8, which has been shown
to express the cardiac isoform exclusively (Donahue et al., 1991;
Zeng et al., 1996
). Neuronal INa was recorded from the murine
neuroblastoma cell line N1E-115. We used reverse transcription
(RT)-PCR analysis to determine the channel isoforms present in
this cell line.
RT-PCR and Sequence Analysis
Total RNA was isolated from N1E-115 cells using the RNeasy Total RNA System (QIAGEN Inc.). RNA was then treated with pancreatic DNase I (Stratagene Inc.) to remove any contaminating
genomic DNA. cDNA was generated by reverse transcription using MuLV Reverse Transcriptase (Perkin-Elmer Corp.) as previously described (Zeng et al., 1996). PCR amplification was carried out using 1.75 U Expand High Fidelity polymerase enzyme
mix (Hoffman-La Roche) with a reaction mixture containing 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 2.25 mM MgCl2, 2% (vol/vol)
DMSO, 0.1% Tween-20, and 1 µM primers as in Dib-Hajj et al.
(1996)
. Five primer sets were selected that would detect various
Na channel
subunits (Table I). Primer sets 1 and 2, which span
a portion of domain IV, S5-S6, are identical to ones used by Dib-Hajj et al. (1996)
. Primer set 1 recognizes isoforms (with Genbank accession numbers) rBrI (X03638), rBrII (X03639), rBrIII
(Y00766), mNa6 (U26707), rPN1 (U79568), mSNS (Y09108),
and rSkm1 (M26643). Primer set 2 recognizes mNav2.3 (L36179), hNav2.1 (M91556), and SCL11 (Y09164). Primer set 3 recognizes sequences upstream of domain I and extends to the
middle of domain I S5 in all rat brain isoforms plus mNa6. A
primer set specific to glyceraldehyde-3-phosphate dehydrogenase
(GAPDH)1 was included as a positive control in each PCR reaction. Although these primers were originally designed based on
rat GAPDH (Genbank X02231), mouse sequence and primers
would be identical except for the 5' base of the reverse primer.
For each primer set, a reaction that contained all of the reaction
components except cDNA (primer only) was also run as a negative control. Using an ABI automatic sequencer, purified PCR-generated fragments were sequenced directly or were first subcloned into the pCRII vector (Invitrogen Corp.).
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Electrophysiology
For whole-cell recordings, the bath solution contained (mM) 140 Na+, 2 Ca2+, 144 Cl, 10 HEPES. To verify accuracy of the measurement of plateau currents, in a few whole-cell experiments
130 Mes
was substituted for 130 Cl
to eliminate an endogenous
Cl
current that had been observed in a few N1E-115 cells; no
differences in Na channel kinetics were observed with this substitution. The pipette solution contained (mM) 140 Cs+, 10 Na+,
100 F
, 50 Cl
, 10 HEPES, set to pH 7.3 with CsOH. Pipette resistances were typically 0.8-1.2 M
. After high-resistance seals were
achieved, cells were lifted from the bottom of the bath and transferred to a chamber containing bath solution supplemented
(when applicable) with saturating concentrations of Anthopleurin B toxin (ApB) and 5 mg/ml bovine serum albumin (to prevent toxin adherence to tubing). Typically, 100 nM toxin was
used with cardiac channels (Kd = 0.9 nM; Khera et al., 1995
),
and 300-600 nM toxin was used with neuronal channels (Kd = 5.1 nM; Khera et al., 1995
) ApB was a kind gift of K. Blumenthal
(University of Cincinnati, Cincinnati, OH). Toxin modification
was monitored by the appearance of prolonged INa in response to
depolarization, and experiments proceeded when modification
was complete. To ensure full channel availability, holding potential was set to
150 (cardiac) or
130 (neuronal) mV.
For single-channel cell-attached recordings, the bath solution
contained (mM) 140 K+, 2 Ca2+, 144 Cl, 10 HEPES, which depolarized the membrane potential to ~0 mV so that pipette potentials did not require correction. The pipette solution contained (mM) 280 Na+, 2 Ca2+, 284 Cl
, 10 HEPES (some early experiments used 140 Na+ and 144 Cl
), set to pH 7.3 with CsOH.
Pipette resistances were typically 2-10 M
, with the lower resistances used for patches exposed to toxin to increase the number
of channels/patch. Pipettes were heat polished, and tips were
painted with Sylgard 182 (Dow Corning Corp.) or sticky wax
(Kerr). We found it difficult to obtain high-resistance seals in the
presence of high toxin concentrations. Therefore, in experiments requiring toxin, the tip of the pipette was filled with toxin-free solution by capillation, and the remainder of the pipette was
back-filled with pipette solution supplemented with 1 µM ApB.
Toxin modification was monitored by the appearance of late
channel reopenings and typically occurred within 2-10 min of establishing a membrane seal.
Voltage protocols were generated using pClamp 6.0.3 (Axon Instruments) and imposed by an Axopatch 200B (Axon Instruments) amplified through a Labmaster TL-1 digital-to-analog convertor. Current waveforms were filtered at 10 (whole cell) or 2 (single channel) kHz, digitized at 50 (whole cell), 25 (single channel, control), or 10 (single channel, toxin) kHz with 12-bit resolution, and stored on an MS-DOS-based computer (Gateway 2000). Temperature was controlled with a TS-4 controller (Sensortek). To resolve single-channel events, the rapid phase of INa recovery, and tail current time course with high accuracy, we carried out all experiments at 6°C. Grouped data are reported as means ± SEM.
Analysis
Whole-cell and single-channel data were analyzed using locally
written software in Matlab 5.1 (The MathWorks). For whole-cell data, traces were ohmically leak corrected based on the current at the holding potential, capacity corrected on the basis of 100 summed subthreshold pulses (20 mV each), and digitally refiltered at 5 kHz using a zero-phase filtering algorithm. Single-channel traces were capacity corrected based on the average of
null sweeps and baseline-corrected by eye. Traces were idealized
using a half-amplitude criterion and checked for accuracy by eye.
For toxin-modified patches, late reopenings were defined as
openings commencing later than 100 ms after depolarization onset and occurring subsequent to a period of at least 15 ms at the
null current amplitude. As in previous single channel studies
with site-3 toxins, occasional subconductance events were noted
(Nagy, 1987; El-Sherif et al., 1992
), and sweeps displaying such
activity were excluded from kinetic analyses.
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RESULTS |
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Na Channel Expression in N1E-115 Cells
The cell line N1E-115 has been a traditional preparation for the evaluation of mammalian neuronal INa
(e.g., Aldrich et al., 1983; Goldman, 1995
), and has also
been studied with respect to site-3 toxins (Khera et al.,
1995
). To characterize the channel isoforms present in
this cell line, we performed RT-PCR experiments using
primer sets complementary to a variety of known Na
channel
subunits. Primer sets 1 and 3 and the positive GAPDH control (Table I) produced the expected
size PCR products (data not shown). No PCR products
were detected using the primer set specific for the
mNav2.3, hNav2.1, or SCL11 sodium channel isoforms
(primer set 2). Except for mNa6, mSNS, and mNav2.3,
full cDNA sequences for most mouse channel isoforms
were not available. For these, we performed sequence
alignments against the rat Na channel
subunits. From
11 subcloned fragments from primer sets 1 and 3, six
sequences were most homologous to rBrII (>90%),
two to rPN1 (>93%), and one to rBrIII (90%). The remaining two subcloned fragments were <68% homologous to any of the reference sequences but were most
similar to rBrI. Excepting rBrI, sequences homologous
to the above listed isoforms were also detected from the
direct sequencing of the PCR-generated fragments.
N1E-115 cells therefore represent an appropriate vehicle for the study of mammalian neuronal INa.
Isoform Dependence of Plateau Current
Consistent with previous results (e.g., Catterall, 1980;
Strichartz et al., 1987
), cardiac and neuronal INa displayed dramatically prolonged decay in the presence of
saturating toxin concentrations. At the conclusion of
this decay, however, a significant plateau current persisted throughout the 2-s test pulses (Fig. 1, A and B).
This current was dependent on voltage and differed in
magnitude with isoform, ranging from 5 ± 1% of the
peak amplitude in cardiac channels to 17 ± 3% of the
peak amplitude in neuronal channels (Fig. 1, C and D).
Despite the differences in magnitude, the voltage dependencies of the plateau currents as evaluated in conductance transforms were similar to that for peak currents (Fig. 1, E and F).
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Steady state voltage-dependent availability was assessed using a uniform protocol with 2-s conditioning
times for both cardiac and neuronal INa. This time was
chosen because it is standard for the cardiac isoform,
and although much longer than is typical in the literature for conditioning neuronal channels, gives similar results to shorter conditioning times (Fozzard and
Hanck, 1996). Availability protocols in the presence of
toxin yielded currents with complex waveforms. In neuronal channels (Fig. 2), test pulses to +20 mV after
negative conditioning steps produced currents with
time courses appropriate to activation of toxin-modified channels from closed states (time to peak, 2.8 ± 0.1 ms). After conditioning at intermediate voltages,
depolarization to the test potential produced a current
that started at zero magnitude and developed monotonically with a time course much slower than that typical of channel activation, reaching a plateau (reflecting the steady state current at Vt = +20 mV) by ~40 ms. After conditioning at positive potentials, current was
present immediately upon shifting to the test potential,
as expected from the presence of the plateau current
during the conditioning pulse and the ohmic change
in driving force. These data were similar to those found
with scorpion toxin in toad axons (Wang and Strichartz, 1985
).
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Depending on the window used for searching for
peak currents and the conditioning potential employed, the steady state availability relationship would
contain varying contributions from all three of these
components: activation, slowly developing plateau current, and already-present plateau current. To calculate
the contribution of that fraction of channels open at
the end of the conditioning pulse (Fig. 2), we measured current at an isochrone immediately after the onset of the test pulse. This relationship was sigmoidal with a half-point and voltage dependence that matched
the conductance relationship of the plateau currents
measured in long depolarizations. Currents measured
at an isochrone corresponding to the time to peak of
INa for activation from a fully available conditioning potential comprised both the instantaneous plateau current measured at the earlier time and the activation of
channels that remained closed and available, although
this time point was still early enough to minimize the
contribution of inactivated channels slowly "recovering" at the test potential of +20 mV. The difference between these two curves, therefore, reflects the "traditional" availability relationship arising from the inactivation of channels during the conditioning pulse. This
relationship and the absence of current immediately after the step to +20 mV from intermediate voltages indicate that channels equilibrate completely into inactivated states at intermediate depolarizations (e.g., 30
mV in Fig. 2 B), but that the inactivated states are incompletely absorbing at more positive voltages.
Cardiac INa displayed an essentially similar, biphasic steady state availability, but the smaller amplitude of the plateau current in this isoform made this behavior more difficult to appreciate. Isochronal steady state availability relations at the time to peak of activation of fully available channels are compared with peak availability relations of control channels for both isoforms in Fig. 3.
Kinetic Basis of Plateau Current
In principle, the plateau current could occur in several
ways. First, the toxin might modify the channels' gating
kinetics such that the inactivated state at positive potential was not completely absorbing, as suggested in a
study in toad myelinated nerve (Wang and Strichartz,
1985). Second, the toxin might induce a (voltage-dependent) fraction of the channels to fail to inactivate completely, yielding a persistent current. Such a modal failure of inactivation has been observed at low frequency
in several mammalian Na channel isoforms (e.g., Patlak
and Ortiz, 1985
; Alzheimer et al., 1993
) and has been
proposed as the mechanism of disease in some human
myotonias (see Cannon, 1996
). It has also been suggested for the mode of action of toxin II from Anemonia
sulcata in skeletal muscle (Cannon and Corey, 1993
).
In short, persistent recovery might occur in all the
channels some of the time, or in some of the channels
all of the time. Finally, toxin binding might affect the
single-channel conductance of modified channels.
Previous single channel studies with site-3 toxins in a
variety of preparations have not reported changes in
single channel conductance (Schreibmayer et al., 1987;
Kirsch et al., 1989
; El-Sherif et al., 1992
; Nagy, 1987
),
but these studies measured currents early after depolarizations. To definitely exclude conductance changes
from playing a role in plateau current, single Na channel events were recorded in the cell-attached patch
configuration in control and with saturating concentrations of toxin added to the pipette solution for both
channel isoforms. The control amplitude was then
compared with that of toxin-modified openings at late times after depolarization onset. Because of the small
amplitude of the plateau current (5-17%), we typically
studied toxin-modified patches with large numbers of
channels. We found that modification by toxin left single-channel conductance unaltered. In [Na]out = 280 mM, conductances were as follows: cardiac control,
17.1 ± 0.6 pS (n = 3); cardiac toxin-modified, 17.6 ± 1.7 pS (n = 4); neuronal control, 23.7 ± 2.7 pS (n = 4);
neuronal toxin-modified, 21.9 ± 1.2 pS (n = 4). It
should be noted that, consistent with other reports
(Nagy, 1987
; El-Sherif et al., 1992
), toxin modification also introduced occasional subconductances, particularly in cardiac channels, which typically occurred at
the beginning or end of excursions to the primary conductance (Fig. 4 A, arrow).
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In the absence of toxin, channels typically inactivated after opening at most a small number of times (Fig. 4, A and C). In channels modified by toxin, however, this behavior was replaced by initial clusters of multiple openings, followed by periodic subsequent reopenings throughout the duration of the recording (Fig. 4, B and D). In accordance with the discrete, stochastic nature of these reopenings, n*Po diaries constructed from recordings of toxin-modified openings over 500-1,000 ms after the onset of depolarizations exhibited moderate activity in a majority of sweeps (Fig. 4, E-H). In contrast, if plateau current was produced through a rare, modal failure of inactivation, n*Po diaries would have contained long trains of openings in only a fraction of sweeps, with accompanying occasional spikes on n*Po diaries. Similar data were obtained in three cardiac patches and three neuronal patches.
We employed two statistical techniques to support
the qualitative observation that persistent current did
not arise through a rare modal failure of inactivation.
First, we employed a runs analysis using the method of
Horn et al. (1984). If even one channel were to fail to
inactivate in any given sweep, the overall value of n*Po
during that sweep would be high. Therefore, we classified sweeps on the basis of aggregate n*Po activity, rating them as either above or below the mean value of
n*Po for that patch and voltage. We then evaluated for
a nonrandom association of high- or low-activity sweeps
using the published statistical criterion (Horn et al.,
1984
). For the 27 data sets (six patches, four voltages for neuronal patches and five voltages for cardiac
patches), only two sets showed any nonrandom activity
(P < 0.05; a N1E-115 patch in two of four voltages
tested,
5 and
25 mV). The remaining voltages of
this patch, as well as the data from the other five patches analyzed in this manner, failed to show any
nonrandom intersweep patterns of activity.
Although the runs analysis failed to support the hypothesis that plateau current arose from modal behavior persistent across multiple sweeps, it did not address
the possibility that the persistent current arose from occasional complete failures of channels to inactivate for
shorter duration; i.e., during single depolarizations. To
test for such patterns for each patch and voltage, we
constructed histograms that grouped sweeps by their
aggregate value of n*Po and fitted these histograms
with a gamma distribution (see, e.g., Johnston and Wu,
1995) (Fig. 4, E, F, and H, insets). Fits were possible for
data sets with >200 openings so that all neuronal data
and two data sets from RT4-B8 patches were analyzed.
Occasional failures of inactivation would be expected to produce very high n*Po values outside the otherwise smooth distribution of activities. To evaluate the extent to which high n*Po sweeps exceeded what would be expected, we identified the tail of the fitted n*Po distribution that accounted for the 10% of the current produced by sweeps with the highest activity. We then compared this with the calculated fraction of actual plateau current produced in this high activity range. Averages for all patches and voltages indicated that 94% of the observed plateau current was explained by periodic, discrete reopenings at late time periods. (Only 6 ± 3% of the plateau current comprised anomalously high n*Po activity.) This analysis and the runs analysis above effectively exclude occasional complete failures of inactivation as a significant contributor to the observed plateau current.
We next compared the mean open times of the late
reopenings to those of control channels. Mean open
times of late reopenings were similar to control open
times at mild depolarizations close to the activation
threshold, and they progressively increased with increasing voltage (Fig. 5). This behavior was similar to that previously recorded from initial, early toxin-modified openings (Kirsch et al., 1989; El-Sherif et al.,
1992
), suggesting that the durations of both early and
late events are influenced by the toxin's inhibition of
inactivation. Even though patches contained multiple
channels, overlapping events at late times were infrequent, as would be expected because of the low probability of channel reopenings. Particularly in cardiac
channels, late re-openings typically comprised of bursts
of several openings with brief closures that were sometimes poorly resolved. To address this possible confounding factor, we constructed histograms measuring
the total open time of bursts (defined as openings separated by closures
3 ms). The mean duration of these
bursts also increased with progressive depolarization
(Fig. 5), implying that the observed voltage dependence of mean open times was not an artifact of missed closures.
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Augmentation of Recovery
The presence of late reopenings separated by long nonconducting periods suggested a model in which the channels' inactivated state(s) were destabilized; i.e., an increase in the rate of recovery from inactivation. We tested this conjecture by measuring whole-cell recovery rates. In both cardiac (Fig. 6) and neuronal (Fig. 7) channel isoforms, modification with ApB significantly enhanced recovery in both isoforms over a range of potentials. This effect was significant at all potentials, as measured by the decrease in the dominant time constant of recovery (Fig. 8, A and C).
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One possible mechanism for this toxin-mediated enhancement might be the enhancement of a recovering
transition directly back to the open state (that is, the I
O transition). Na channels do not normally recover
via this path (Kuo and Bean, 1994
). However, the demonstrated ability of these toxins to inhibit the O
I transition gives this prospect a pleasing symmetry. During recovery, inactivated Na channels must normally
move through several inactivated states before reaching
a state with an appreciable rate of return into the activation pathway (Scheme I; Chiu, 1977
; Kuo and Bean, 1994
). Thus, recovery proceeds after an initial delay, reflecting the time required for the channel's transit
through one or more inactivated states. This delay was
easily detectable for both channel isoforms in control
at the experimental temperature of 6°C (Figs. 6 and 7,
insets). Previous data noted an absence of this delay in
toad myelinated nerve after channel modification (Wang and Strichartz, 1985
). Also, because of the existence of a toxin-induced plateau current that persisted
until the end of the inactivating prepulses, we expected
some INa even at very short recovery times. Therefore,
we were surprised to find that a delay persisted in toxin
before any channels, in addition to those already comprising the plateau current, recovered into the activation pathway. This delay was shorter than that found in control channels and became undetectable only at the
most negative recovery voltages tested. This lag is
graphically evident in Figs. 6 B and 7 B (arrows) and is
quantified in Fig. 8, B and D.
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The continued presence of a delay even when a fraction of channels was open at the onset of the recovery
interval strongly argues against significant recovery of
channels through the open state at negative potentials.
To further verify that toxin did not augment recovery
directly into the open state, we recorded current during the recovery process. That is, cells expressing cardiac or neuronal channels were depolarized until inactivation reached steady state and were allowed to recover while current was recorded at high resolution.
The protocol is depicted in Fig. 9 A. In cells expressing
neuronal channels (Fig. 9 B), a current was present at
the onset of the recovery interval and it rapidly decayed. We interpret this current as arising from channels open at the end of the conditioning prepulse conducting current according to the increased driving
force at the recovery potential. The time constant of
this current decay (290 µs at 100 mV in Fig. 9 B) was
much faster than the time constant of recovery from inactivation (13.6 ms at
100 mV, from Fig. 8 C) and was
similar to the time constant of deactivation of unmodified INa tail currents at
100 mV for the same cell
shown in Fig. 9 B (240 µs, data not shown). The behavior of cardiac channels was qualitatively similar (Fig. 9
C), but the smaller magnitude of the plateau current in
this isoform made the detection of current during recovery more difficult. In neither isoform was a discrete
current arising from recovering channels detectable
(see DISCUSSION).
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DISCUSSION |
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In this study, we have demonstrated the existence of a
noninactivating plateau current in both cardiac and
neuronal isoforms of the voltage-gated Na channel
when modified by a high affinity site-3 sea anemone
toxin (Anthopleurin B). The voltage dependence of
the equilibrium distribution between nonconducting
and conducting states is biphasic. That is, toxin-modified channels reside entirely in closed states at negative
potentials (as do toxin-free channels), partition completely into inactivated states at intermediate voltages, and occupy the open state with an appreciable probability at positive potentials. In this sense, our results accord strongly with those obtained from channels modified with -scorpion toxins (Koppenhofer and Schmidt,
1968
; Mozhayeva et al., 1980
; Wang and Strichartz, 1985
), where the steady state current relation also
showed a nadir at modest depolarizations. At first
glance, these results appear to conflict with a number
of other reports of toxin-modified channel availability.
An early study in the frog node of Ranvier with toxin II
from A. sulcata (Bergman et al., 1976
) produced an
availability curve that reached zero, but the toxin concentrations used were near the Kd for the toxin/channels studied, probably leading to an underestimation of
the plateau current magnitude. A study of the action of
Leiurus toxin on N18 neuroblastoma cells also demonstrated only a small plateau current, but these investigators did not probe the relationship at positive potentials (Gonoi et al., 1984
). Likewise, the apparent complete inactivation noted in canine cardiac myocytes
(Hanck and Sheets, 1995
) undoubtedly arose because
of the low amplitude of the plateau current in that
channel isoform and the relatively negative voltage
range tested. On the other hand, the voltage-independent plateau current noted by Wasserstrom et al.
(1993)
probably reflects the steady state current at
their test potential (
20 mV).
Analysis of single channel recordings revealed that
the plateau current arises from discrete, late channel
reopenings. These late events have kinetics similar to
early toxin-modified events and are separated by longer
closed intervals, and so appear to reflect an increase in
one or more of the rate constants in the recovery pathway, a hypothesis also supported by the toxin-mediated enhancement of whole-cell recovery from inactivation.
Consistent with these observations, two previous single-channel studies reported that toxin treatment induced
a second, slow component into the first-latency distribution of channel openings; that is, a subset of sweeps
showed activity in which channels first opened at dramatically late times after depolarization onset (Schreibmayer et al., 1987; Kirsch et al., 1989
). These papers
did not consider the issue of late reopenings; however,
in light of these studies, this late component may represent channels that inactivate without opening upon depolarization onset, but later recover back into the activation pathway and open for the first time.
Site-3 toxins do not induce a modal shift in gating, as
evidenced by the relative homogeneity of the n*Po diaries and failure of statistical analysis to detect modal activity across or within sweeps. The suggestion of this action by toxin II from A. sulcata in skeletal muscle Na
channels (Cannon and Corey, 1993) was probably a result of the low affinity for this toxin for skeletal muscle channels, causing patches to be modified during only a
fraction of the sweeps.
These toxins are well known to have lower affinity for
inactivated than for resting or open channels (Catterall, 1979; Mozhayeva et al., 1980
; Warashina and Fujita,
1983
; Strichartz and Wang, 1986
). In fact, for some
toxin/channel pairs, this can be dramatic and easily observed (Strichartz and Wang, 1986
). Consequently, we
considered whether dissociation of toxin during prolonged depolarization might occur. However, this process was minimal in these experiments. First, the Na+
current induced by the toxins included a persistent current that did not decay appreciably during a 1.5-s depolarization (see Fig. 1, A and B), a time comparable to
that of the longest conditioning pulse used in these
studies (2 s). By definition, therefore, toxin did not dissociate significantly during this time interval. Second, if
toxin were to dissociate during extended conditioning
pulses, upon repolarization it could not rebind to channels at a rate any faster than that dictated by the known
toxin/channel association rates, which are slow (Khera
et al., 1995
). For instance, N1E-115 channels in the
presence of 300 nM ApB (a commonly used concentration) would be expected to rebind toxin with a time
constant of 3.5 s, but after a depolarization channels
were fully modified at the earliest times channels recover (e.g., 0.25 s in Fig. 7). Finally, we directly assayed
toxin dissociation by depolarizing toxin-modified neuronal cells to 0 mV in toxin-free solution for 20 and 60 s,
and only 5.4 and 11.5% dissociation occurred. It is,
therefore, unlikely that the dissociation of toxin from
channels during depolarization affects the results.
Although there appear at first glance to be differences in the effect of toxin on INa from cardiac and neuronal channels (compare, for example, Fig. 1, A and B), these differences are most likely to be secondary to quantitative differences in the kinetics of the two isoforms. In both isoforms, toxin binding appears to augment the rate of recovery (compare Fig. 8, A and C), shorten but not eliminate the delay before recovery onset (compare Fig. 8, B and D), and produce a plateau current comprising discrete late reopenings (compare Fig. 4, B and D) with no evidence for modal inactivation failure. The relatively prolonged mean open and burst times found in cardiac channels as compared with neuronal channels (Fig. 5) probably reflect only differing rate constants in the activation pathway. Taken together, modification by toxin in cardiac channels produces a plateau current of smaller magnitude, comprising fewer, longer openings than in neuronal cells, but arising through the same enhancement of recovery from inactivation as found in neuronal channels.
Analysis of recovery in whole-cell experiments demonstrated that toxin speeds recovery at all potentials
and shortens the delay in recovery from inactivation. In
this regard, our data stand in contrast to those reported
in toad myelinated nerve since in that preparation
toxin eliminated the delay before the onset of recovery
and did not change the time constant of current recovery (Wang and Strichartz, 1985).
How might toxin treatment destabilize the channels'
inactivated states and enhance macroscopic recovery?
The accepted existence of a delay before the onset of
recovery in Na channels (Kuo and Bean, 1994) strongly
suggests the presence of multiple inactivated states. In
light of this observation, several kinetic modifications might produce the observed enhancement in recovery.
First, toxins might augment the rate of recovery into
the open state (Scheme II, 1). Second, toxins might enhance one or more of the existing rates of channel recovery to closed, available states (Scheme II, 2). Third,
toxins might simply enhance one or more deactivating (i.e., "leftward") transition rates between the multiple
inactivated states, bringing channels more quickly into
inactivated states from which recovery rates are high
(Scheme II, 3). Finally, the action of toxin might of
course comprise a combination of these effects. Site-3
toxins are already known to decrease the rate of Na
channel inactivation from the open state, O
I
(Schreibmayer et al., 1987
; Nagy, 1988
; Kirsch et al.,
1989
; El-Sherif et al., 1992
; Hanck and Sheets, 1995
;
Sheets and Hanck, 1995
). Therefore, it is tempting to
hypothesize that the plateau current at positive voltages
represents an O
I equilibration that partially favors the open state, and we can predict quantitative boundaries on the unidirectional rate constants of O
I and
I
O.
|
The rate of decay of macroscopic Na+ current after
toxin modification is remarkably voltage independent.
In neuronal channels treated with saturating ApB, single exponential fits to the decaying phase of current
transients yielded a time constant of 31.4 ± 0.4 ms, invariant over a range of 25 to +30 mV (n = 5). At the positive end of this voltage range, the majority of channels are likely to be found in either the open state or
the distal-most inactivated state. The reciprocal of this
time constant can, therefore, be taken as representing
the rate of equilibration between these two conformations (kOI + kIO). The O
I equilibrium places 17% (plateau amplitude at
+20 mV in Fig. 2) of the channels in the open state [kIO/(kOI + kIO)]. Thus, kOI is calculated to be 26 s
1, and kIO is calculated as 5 s
1. This
rate of I
O transitions is significantly slower than the
observed macroscopic rate of recovery from inactivation in toxin-modified N1E-115 cells (see Fig. 8 C). Even
in the presence of toxin, therefore, channels at negative voltages more likely recover by traversing multiple
inactivated states before returning to one of the closed
states. Consistent with this conclusion, a delay in recovery from inactivation continues to be measurable except at the most negative voltages (see Fig. 8, B and D).
The issue of channel recovery through the open state
during repolarization can also be addressed by the time
course of current present during recovery from inactivation. We can evaluate the contribution of kIO by simulation using an irreversible, simplified Markov model of
the recovery process (Fig. 10), using rate constants based on the experimental data from neuronal channels. The closing rate (kOC) was taken from the time
constant of tail currents in the absence of toxin (at
100 mV), and the rate of recovery (kIC) was taken
from the average rate of recovery at
100 mV in the absence of toxin. The form of the current during recovery depends on the relative magnitude of the rate constant between I and O (kIO) compared with kOC and kIC.
If kIO is similar to the rate of deactivation (kOC), a
"hooked" current will appear during recovery, reflecting the accumulation of channels in O (e.g., kIO = 1,500). If kIO is slower than kOC but still faster than kIC,
then current will decay with an apparent exponential,
but the time constant of this exponential will approximate the overall time constant of recovery (e.g., kIO = 500). If kIO is slow, but still sufficiently large to cause appreciable recovery through the open state (e.g., kIO = 77), then current decay will comprise two obvious components: a rapid deactivation and a much slower decay
corresponding to recovering channels.
|
The observed behavior (e.g., Fig. 9 B, 100 mV)
shows a complete exponential decay of current that is
rapid in comparison to the rate of whole-cell recovery,
which is similar to that expected when kIO is low (e.g.,
Fig. 10, 18 s
1) or nonexistent (Fig. 10, dotted line).
In summary, therefore, it appears that the plateau
current produced in the presence of toxin arises from
an O I equilibrium that partially favors the open
state, but that the overall rate of I
O recovery is still
not sufficiently large to cause appreciable numbers of
channels to recover through the open state during repolarization. Nonetheless, toxin treatment does augment recovery from inactivation. We suggest that both
of these observations can be parsimoniously rationalized under the assumption that the toxin destabilizes the terminal inactivated state of the channel. Open-state inactivation is slowed (Schreibmayer et al., 1987
;
Nagy, 1988
; Kirsch et al., 1989
; El-Sherif et al., 1992
),
producing the well-known prolongation of macroscopic inactivation and increase in mean open time.
This slowing, combined with a possible augmentation
of the open-state recovery rate I
O, produces the observed plateau current. Finally, destabilization of the final inactivated state augments the leftward I
I transition, enhancing recovery and shortening the delay before recovery onset.
This hypothesis accounts for all of the observed kinetic features of this toxin through destabilization of a single kinetic state of the channel, raising the possibility that this toxin can be used to define the molecular determinants of a specific kinetic state. Such a molecular identification has not been previously possible, and it would be an important step towards reconciling kinetic models with gross channel conformation.
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FOOTNOTES |
---|
Address correspondence to Dr. Dorothy Hanck, MC6094, The University of Chicago, 5841 S. Maryland Ave., Chicago, IL 60637. Fax: 773-702-6789; E-mail: d-hanck{at}uchicago.edu
Original version received 8 September 1998 and accepted version received 10 December 1998.
Portions of this work were previously published in abstract form (Benzinger, G.R., and D.A. Hanck. 1997. Biophys. J. 72:A362).We thank Kenneth Blumenthal for providing the recombinant ApB toxin. We thank Jack Kyle and Ruth Martin for their on-going contributions to our research, and Michael Sheets (University of Utah, Salt Lake City, UT) for his critical reading of the manuscript.
This study was supported by National Heart, Lung and Blood Institute grant HL-PO1-20592 (D.A. Hanck) and an American Heart Association Established Investigatorship (D.A. Hanck). G.R. Benzinger is a trainee of the Medical Scientist Training Program (5-T32-GM07281).
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Abbreviations used in this paper |
---|
ApB, Anthopleurin B toxin; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; RT-PCR, reverse transcription-PCR.
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