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The Journal of Neuroscience, March 15, 2001, 21(6):1884-1892
Recovery from Inactivation of T-Type
Ca2+ Channels in Rat Thalamic Neurons
Chung-Chin
Kuo1, 2 and
Shibing
Yang1
1 Department of Physiology, National Taiwan University
College of Medicine, and 2 Department of Neurology,
National Taiwan University Hospital, Taipei 100, Taiwan
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ABSTRACT |
We studied the gating kinetics, especially the kinetics of
recovery from inactivation, of T-type Ca2+ channels
(T-channels) in thalamic neurons. The recovery course is associated
with no discernible Ca2+ current and is
characterized by an initial delay, as well as a subsequent exponential
phase. These findings are qualitatively similar to previous
observations on neuronal Na+ channels and suggest
that T-channels also must deactivate to recover from inactivation. In
contrast to Na+ channels in which both the delay and
the time constant of the exponential phase are shortened with
increasing hyperpolarization, in T-channels the time constant of the
exponential recovery phase remains unchanged between  100 and 200
mV, although the initial delay is still shortened e-fold
per 43 mV hyperpolarization over the same voltage range. The
deactivating kinetics of tail T-currents also show a similar voltage
dependence between 90 and 170 mV. According to the hinged-lid model
of fast inactivation, these findings suggest that the affinity
difference between inactivating peptide binding to the activated
channel and binding to the fully deactivated channel is much smaller in
T-channels than in Na+ channels. Moreover, the
inactivating peptide in T-channels seems to have much slower binding
and unbinding kinetics, and the unbinding rates probably remain
unchanged once the inactivated T-channel has gone through the initial
steps of deactivation and "closes" the pore (with the activation
gate). T-channels thus might have a more rigid hinge and a more abrupt
conformational change in the inactivation machinery associated with
opening and closing of the pore.
Key words:
T-type Ca2+ channel; activation; deactivation; inactivation; recovery from inactivation; gating
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INTRODUCTION |
Fast inactivation is an important gating phenomenon in many
voltage-gated ion channels, such as Na+,
Shaker K+, and T-type
Ca2+ channels (T-channels). Upon membrane
depolarization, these channels are rapidly activated and then rapidly
inactivated. Fast inactivation thus tightly controls ion flow through
the channel. A classical mechanistic view of fast
inactivation is the "ball-and-chain" model (Armstrong and
Bezanilla, 1977 ; Armstrong, 1981), in which inactivation results from
blockade of the activated channel pore by part of the channel protein
(such as the N terminal region in Shaker
K+ channels) (Hoshi et al., 1990; Zagotta
et al., 1990). More recently, West et al. (1992) proposed a somewhat
different "hinged-lid" model, in which the linker peptide between
transmembrane domains III and IV of the
Na+ channel protein functions as a
"lid" to control ion permeation at the internal pore mouth. Both
models, however, are the same in postulating fast inactivation as
open-channel blockade produced by binding of the inactivating peptide
to a receptor uncovered or produced by channel
activation.
Scheme 1 is a simplified diagram incorporating the foregoing
concepts of fast inactivation, in which OB and CB denote the open
(activated) and closed (deactivated) conformations blocked by the
inactivating peptide, respectively. In principle, route C to O to OB is
the major (or even exclusive) pathway for the development of
inactivation, so that inactivation is coupled to activation. On the
other hand, the inactivated channel (state OB) may recover through the
OB to O to C (unblocking-first) route or the OB to CB to C
(deactivation-first) route. The unblocking-first route implies
substantial ionic current through the channels traversing state O
during recovery. The deactivation-first route ensures no such currents,
but the recovery time course might be characterized by a
voltage-dependent initial delay that corresponds to deactivation of the
inactivated channels (OB to CB step). Inactivated
Na+ channels always take the
deactivation-first route to recover (Kuo and Bean, 1994), whereas
recovery of inactivated Shaker K+ channels
favors the OB to O to C route (Kuo, 1997).
The molecular mechanisms underlying the development and recovery of
fast inactivation in T-channels have been less extensively studied. The
macroscopic inactivation rates of T-channels saturate at positive
potentials (Chen and Hess, 1990; Herrington and Lingle, 1992; Serrano
et al., 1999), indicating lack of intrinsic voltage dependence of the
inactivation process itself. There is no obvious Ca2+ current associated with recovery of
cloned ( 1G) T-channels at 100 mV (Serrano et al., 1999).
Also, there is an initial delay in the recovery time course of the
cloned 1G and 1H channels at 100 mV (Satin and Cribbs, 2000).
These findings would support the deactivation-first route of recovery.
However, more rigorous examination of the initial delay and the current
associated with recovery at different recovery potentials seems
necessary, because the choice of recovery route may be different at
different potentials (Kuo, 1997). The recovery kinetics of fibroblast
or cloned (1G) T-channels are voltage-independent between 100 to
130 mV but become slower at more positive potentials (Chen and Hess,
1990; Serrano et al., 1999). However, the recovery kinetics of
T-channels in rat GH3 pituitary cells remain unchanged throughout 80
to 130 mV (Herrington and Lingle, 1992), suggesting different
biophysical properties of T-channels in different tissues. We therefore
studied the recovery of inactivated T-channels in thalamic neurons, in which T-channels play an essential role in controlling the discharge pattern (for review, see McCormick and Bal, 1997). We found that, like
Na+ channels, T-channels also take the
deactivation-first route to recover from inactivation. However, the
differential work it takes to deactivate the inactivated (state OB) and
the open (state O) channels seems to be much smaller in T-channels than
in Na+ channels. Also, T-channels probably
have a more rigid hinge and a more abrupt conformational change in the
inactivation machinery associated with opening and closing of the pore.
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MATERIALS AND METHODS |
Cell preparation. Coronal slices of the whole brain
were prepared from 8- to 12-d-old Wistar rats. The brain was cooled in 0°C cutting solution (125 mM sucrose, 20 mM
Na2SO4, 10 mM
K2SO4, 3 mM MgCl2, and 10 mM HEPES, pH 7.4) for 1 min. The tissue block containing thalamus was then cut into 400 µm slices with a vibratome (Campden Instruments, Silbey, UK). The ventroposterior complex (including the ventroposterior medial and ventroposterior lateral nuclei) of thalamus was dissected from the slices and cut into small
pieces. After treatment for 3-5 min at 33°C in dissociation medium
(82 mM
Na2SO4, 30 mM
K2SO4, 0.4 mM CaCl2, 3 mM MgCl2, 5 mM HEPES, and 0.001% phenol red indicator, pH
7.4) containing 0.5 mg/ml trypsin (type XI; Sigma, St. Louis, MO), the
tissue pieces were moved to dissociation medium containing no trypsin but 2 mg/ml bovine serum albumin (Sigma). Each time when cells were
needed, two to three pieces were picked and triturated to release
single neurons.
Whole-cell recording. The dissociated neurons were put in a
recording chamber containing Tyrode's solution (150 mM NaCl, 4 mM KCl, 2 mM MgCl2, 2 mM CaCl2, and 10 mM HEPES, pH 7.4). Seal was formed, and
whole-cell configuration was obtained in Tyrode's solution. The cell
was then lifted from bottom of the chamber and moved in front of an
array of flow pipes (content 1 µl, length 64 mm; Microcapillary;
Hilgenberg Inc., Malsfeld, Germany) emitting different external
solutions. The external solution for recording Ca2+ currents contains 150 mM tetraethylammonium chloride, 5 mM CaCl2, 10 mM HEPES, and 3 µM
tetrodotoxin, pH 7.4. Except for the experiments in Figure
1A-D, 1 µM nimodipine and
0.5 µM  -conotoxin MVIIC were always added to
the external solution to prevent contamination of T-current by L-, N-,
and P/Q-type Ca2+ currents. Whole-cell
voltage-clamp recordings were made using pipettes pulled from
borosilicate micropipettes (outer diameter, 1.55-1.60 mm;
Hilgenberg Inc.), fire polished, and coated with Sylgard (Dow Corning,
Midland, MI). The standard internal solution contained 90 mM
N-methyl-D-glucamine fluoride, 45 mM
N-methyl-D-glucamine chloride, 7.5 mM EGTA, 1.8 mM
MgCl2, 9 mM HEPES, 4 mM MgATP, 0.3 mM GTP (Tris
salt), and 14 mM creatinine phosphate (Tris
salt), pH 7.4. The intracellular fluoride ion may help to abolish the L-type Ca2+ current (Akaike et al., 1983;
Carbone and Lux, 1987). The capacitance and series resistance of the
whole-cell configuration were mostly 5-20 pF and 2-5 M (after a
typical 40-60% partial compensation), respectively. Currents were
recorded at room temperature (~25°C) with an Axoclamp 200A
amplifier, filtered at 2-10 kHz with four-pole Bessel filter,
digitized at 20-200 µsec intervals, and stored using a Digidata-1200
analog-to-digital interface along with the pClamp software (Axon
Instruments, Foster City, CA). All statistics are given as mean ± SEM.
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RESULTS |
Block of the low-voltage-activated Ca2+ current
by micromolar La3+
Figure 1 shows typical
Ca2+ currents recorded from neurons in the
ventroposterior nuclei of thalamus. In Figure 1, A and
B, the cell contains a significant amount of sustained
Ca2+ currents, which is much more manifest
with a test pulse of 10 mV than of 40 or 70 mV (i.e.,
high-voltage-activated) and is not very sensitive to a holding
potential change from 120 to 80 mV. In contrast, the
Ca2+ currents in the other cell (Fig.
1C-E) consist almost exclusively of transient currents. The
transient currents are significantly activated with test pulses 70 to
40 mV (low-voltage-activated) and are rapidly inactivated with a
decaying time constant of 20-25 msec. These transient currents are
insensitive to 1 µM nimodipine or 0.5 µM -conotoxin MVIIC, yet are very sensitive
to more depolarized holding potentials. Little transient
Ca2+ current can be recorded from the same
cell when the holding potential is changed from 120 to 80 mV. The
rapid inactivation, the low-voltage activation, and the sensitivity to
holding potential all suggest that the chief component of these
transient neuronal Ca2+ currents is
contributed by the T-type Ca2+ channel
(T-channels) (Carbone and Lux, 1987; Fox et al., 1987; Coulter et al.,
1989). This latter cell thus is classified as type II neurons (Fig. 1,
legend), which contain almost only T-channels and have negligible
high-voltage-activated Ca2+ currents. The
former cell containing significant high-voltage-activated Ca2+ currents is classified as type I
neurons. Among the 52 cells we examined, 65% (34 of 52) are type II
and 35% (18 of 52) are type I. All of the subsequent studies are
performed in type II neurons.
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Figure 1.
Two types of thalamic relay neurons with
different compositions of whole-cell Ca2+ currents.
Nimodipine and -conotoxin MVIIC were not added in
A-D but were added in E.
A, The cell was held at 120 mV and stepped every 5 sec
to the test pulse (100, 70, 40, and 10 mV) for 200 msec. A test pulse of 100 mV
elicited no Ca2+ currents, which started to be
discernible with a test pulse of 70 mV. At 40 mV, the
Ca2+ currents were predominated by a rapidly
inactivating or transient component (presumably T-current), whereas a
more sustained component (presumably L- or N-type or other
high-voltage-activated Ca2+ currents) became very
manifest at 10 mV. B, The same cell and pulse protocol
as that in A, except that the holding potential was
changed to 80 mV. Note a significant decrease of the transient
component but not the sustained component of the
Ca2+ currents. C, The same pulse
protocol as that in A (holding potential of 120 mV)
was repeated in another cell. There was no current elicited by a test
pulse of 100 mV, but significant transient current was observed with
a test pulse of 70 mV. Most interestingly, no sustained current was
observed with test pulses of 40 or even 10 mV. D,
The same cell and pulse protocol as that in C, except
that the holding potential was changed to 80 mV. No current was
elicited with test pulses of 100 to 10 mV. E, The
same cell and pulse protocol as that in C, except that
0.5 µM -conotoxin MVIIC and 1 µM
nimodipine were added to the external solution. Note that the transient
Ca2+ currents are slightly decreased in amplitude,
but kinetics of the currents and the current-voltage relationship
remain unchanged. F, The cells in which
Ca2+ current is larger at a test pulse of 10 mV
than at a test pulse of 40 mV are classified as type I neurons
(typically represented by the cell in A). The cells in
which Ca2+ current is smaller at a test pulse of
10 mV than at a test pulse of 40 mV are classified as type II
neurons (typically represented by the cell in C), which
always contain little discernible high-voltage-activated
Ca2+ currents.
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Like in the other Ca2+ channels,
La3+ is a potent blocker of T-channels, in
which the dissociation constants are ~0.5 µM (in dorsal root ganglion cells; Todorovic and Lingle, 1998) and 2.5 µM (in clonal GH3 pituitary cells; Herrington and Lingle,
1992). Figure 2A shows
that most of the rapidly inactivating current in thalamic neurons is
blocked by 10 µM
La3+. The difference between the currents
recorded in the presence and absence of 10 µM
La3+ is the
La3+-sensitive current. This differential
current will be used later in the other experiments when it is
necessary to eliminate capacity transient and tiny contamination of any
nonspecific currents.
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Figure 2.
La3+ blockade of T-current.
A, The cell was held at 120 mV and stepped every 5 sec
to 50 mV for 300 msec to elicit T-current. The same pulse protocol
was repeated in the presence or absence of 10 µM external
La3+. It is evident that 10 µM
La3+ inhibits most of the T-current.
B, The La3+-sensitive current is
obtained by subtracting the current in the presence of external
La3+ from the current in the control external
solution.
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Saturation of the macroscopic inactivation rate at voltages more
positive than 40 mV
Figure 3 plots the peak T-current
and the decaying time constant against time. The current size and the
inactivation kinetics do not show significant changes in 30 min, which
is long enough for collecting most data in this study. Figure
4A-C examines the voltage dependence of activation as well as inactivation kinetics and
the steady-state inactivation curve of T-channels. The activation phase
is speeded by increasing depolarization between 80 and 10 mV. On
the other hand, the inactivation kinetics are accelerated with
increasing depolarization only between 80 and 40 mV, yet become
saturated at more positive voltages. In terms of Scheme 1, it
seems that the horizontal transitions (e.g., C to O) are voltage-dependent, yet the vertical transitions (e.g., O to OB) are
voltage-independent. This is similar to the case of
Na+ channels (Bezanilla and Armstrong,
1977; Bean, 1981; Aldrich et al., 1983; Gonoi and Hille, 1987). Fast
inactivation of T-channels thus probably takes the C to O to OB route
to develop and gets its apparent voltage dependence from the voltage
dependence of activation. The saturated inactivation time constant
(~20 msec) (Fig. 4B) could indicate a
voltage-independent O to OB rate of 50 sec 1. Figure
5A-C demonstrates
deactivating tail currents of T-channels. The deactivation kinetics
(i.e., rates of the O to C transition) are accelerated exponentially
from 90 to 170 mV, with an apparent voltage dependence of
e-fold acceleration per ~30 mV hyperpolarization.
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Figure 3.
Insignificant rundown of T-current in 30 min.
A, The cells were held at 120 mV, and T-current was
elicited by the same pulse protocol as that in Figure 2. The peak
T-currents obtained at each time point are normalized to the peak
T-current of the first sweep in each cell. The mean normalized peak
currents from four cells are then plotted against time. There is no
significant decrease of the peak T-current in a period of 30 min.
B, The decaying phase of the T-current in
A is fitted by monoexponential functions, and the
decaying time constant is plotted against time. The time constants show
no significant change in 30 min.
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Figure 4.
Voltage dependence of macroscopic activation and
inactivation of T-current. A, The cells were held at
120 mV and stepped every 5 sec to 10 to 80 mV for 500 msec to
elicit T-current. La3+-sensitive currents were used
to eliminate capacity transient and facilitate analysis of the
currents. The rising phase of the macroscopic currents (from the
beginning to the peak of each current) are fitted by monoexponential
functions. The time constants from the fits are 5.6 ± 1.0, 3.2 ± 0.2, 1.9 ± 0.2, 1.3 ± 0.1, 0.89 ± 0.11, 0.61 ± 0.08, 0.48 ± 0.1, and 0.40 ± 0.07 msec for
step potentials 80, 70, 60, 50, 40, 30, 20, and 10 mV,
respectively (all n = 4; note that the
vertical axis is in logarithmic scale).
B, The decaying phase of the current sweeps in
A are also fitted by monoexponential functions, and the
time constants are 43.9 ± 8.0, 24.9 ± 4.2, 18.8 ± 2.3, 16.7 ± 1.7, 15.5 ± 1.2, 15.0 ± 0.8, 14.8 ± 0.8, and 15.7 ± 0.9 msec for step potentials 80, 70, 60,
50, 40, 30, 20, and 10 mV, respectively. C, A
representative inactivation curve of T-channels from four type II
thalamic neurons. The cells are held at 120 mV and stepped every 5 sec to the inactivating pulse (50 to 150 mV) for 500 msec. The
channels that remained available after each inactivating pulse were
assessed by the peak currents during a following short pulse to 50 mV
for 500 msec. The fraction available is defined as the normalized peak
current (relative to the peak current evoked with an inactivating pulse
at 150 mV) and is plotted against the voltage of the inactivating
pulse. The line is a fit to the data with a Boltzmann function:
fraction available = 1/(1 + exp((V + 96.8)/7.3)),
where V denotes the voltage of the inactivating pulse in
millivolts.
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Figure 5.
Voltage dependence of macroscopic deactivation of
T-current. A, The cell was held at 120 mV and stepped
every 5 sec to 50 mV for 5 msec (the activating pulse) and then to
90 to 170 mV for 30 msec (the deactivating pulse).
La3+-subtracted currents were used to eliminate
capacity transients and facilitate subsequent fitting procedures. The
tail currents show larger amplitude and faster decaying kinetics as the
deactivating pulse goes more negative. B, The decaying
phase of tail current is fitted by monoexponential functions. The
decaying time constants are 6.1 ± 1.5, 4.6 ± 0.7, 3.2 ± 0.4, 2.2 ± 0.3, 1.6 ± 0.2, 1.1 ± 0.1, 0.81 ± 0.07, 0.61 ± 0.06, and 0.47 ± 0.04 msec for deactivating
potentials 90, 100, 110, 120, 130, 140, 150, 160, and
170 mV, respectively (all n = 5).
C, The mean value of time constant in B
is plotted against the voltage of the deactivating pulse in a
semilogarithm scale (the longitudinal axis is the
natural logarithm of the deactivating time constant in milliseconds).
The line is a linear fit of the form: ln( ) = 4.77 + 0.033V, where V denotes voltage of
the deactivating pulse in millivolts.
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No Ca2+ currents associated with recovery
from inactivation
If the inactivated T-channels take the OB to O to C
(unblocking-first) route to recover from inactivation, there might be significant Ca2+ current produced by
T-channels traversing the O state during the hyperpolarizing recovery
pulse. On the other hand, if the inactivated T-channels take the OB to
CB to C (deactivation-first) route, there would be no such
Ca2+ current. Figure
6 shows that the latter is true not only
with mildly (80 mV) but also with more strongly (120 mV)
hyperpolarizing recovery pulses. After most T-channels are inactivated
by a pulse to 40 mV, there is no discernible current during the
following hyperpolarization phase to either 80 or 120 mV. This
finding suggests that inactivated T-channels take the
deactivation-first route to recover, regardless of the recovery
potential.
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Figure 6.
Lack of discernible current during the recovery
phase of inactivated T-channels. The cell was held at 120 mV and
stepped to 40 mV for 500 msec (the inactivating pulse) and then
stepped back to 80 mV (top) or 120 mV
(bottom) to recover the inactivated T-channels.
La3+-subtracted currents are used to eliminate
contamination from leak or other nonspecific currents. No matter that
the recovery potential is 80 or 120 mV, there is no discernible
ionic current associated with recovery of the inactivated
T-channels.
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Gradual shortening of the initial delay with
increasing hyperpolarization
If inactivated T-channels take the deactivation-first route to
recover, then there may be an initial delay corresponding to the OB to
CB step in the time course of recovery. Figure
7, A and B,
shows that there is indeed an initial delay in the
recovery course of inactivated T-channels in thalamic neurons.
Moreover, the delay gets shorter with increasing hyperpolarization of
the recovery pulse (down to 200 mV), showing an apparent voltage dependence of e-fold shortening per 43 mV hyperpolarization
(Fig. 7C). The voltage dependence of the delay suggests
involvement of charge movements in the pathway leading to the channel
conformation from which the inactivating peptide unbinds. If the
channel increases its affinity for the inactivating peptide as a result
of the forward movement of the voltage sensors (i.e., channel
activation), it is likely that backward movements of the sensors, or
deactivation, would decrease the affinity of the blocking peptide and
facilitate the unbinding process. This is qualitatively similar to the
case of Na+ channels (Kuo and Bean, 1994)
and is consistent with the view that the initial delay results from
deactivation of the inactivated channels. The initial delay in
T-channels, however, is quantitatively very different from that in
Na+ channels, especially when taking the
deactivating kinetics into consideration (see Discussion).
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Figure 7.
Initial delay in the recovery time course of
T-channels over a wide range of membrane potentials. A,
The cells was held at 120 mV and pulsed twice to 50 mV (each for
200 msec) every 6 sec, with a gradually lengthened gap between the two
pulses at various potentials (the recovery potential,
Vr). The average current in the last
2 msec in the first pulse is subtracted from the peak currents in both
pulses to make "corrected" peak currents. The fraction of recovered
channels is determined by the ratio between the corrected peak current
in the second pulse and that in the first pulse. Because the fraction
of recovered channels with such short Vr is
very small, we try to eliminate the contamination from noises by a
smoothing technique. The average value of three consecutive data points
is calculated and designated as the final value of the middle point of
the three and is plotted against the duration of
Vr. It is evident that there is an initial
delay in the recovery course with a Vr of
120 mV. The delay becomes shorter with more hyperpolarized
Vr and is almost negligible with a
Vr of 200 mV. Also note that the recovery
courses immediately after the delay are approximately linear and are of
almost identical slope with Vr of 120,
160, or 200 mV. This is consistent with the findings in Figure 8
(see below), which shows that the recovery course after the initial
delay can be approximated by monoexponential functions with almost
identical time constants at different Vr.
B, With the analysis given in A, if one
data point is larger than its previous point, the interval between the
two points is defined as an "increment." The initial delay is
defined by the first data point that marks the start of four
consecutive increments. The initial delays from seven to nine cells are
13.5 ± 2.5, 8.2 ± 1.3, 4.6 ± 0.6, 3.1 ± 0.5,
2.1 ± 0.4, and 1.3 ± 0.3 msec for Vr of
100, 120, 140, 160, 180, and 200 mV, respectively.
C, The mean value of the initial delay in
B is plotted against Vr in a
semilogarithm scale (the longitudinal axis is the
natural logarithm of the initial delay in milliseconds). The line is a
linear fit of the form: ln(delay) = 4.86 + 0.023V, where V denotes
Vr in millivolts.
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Saturation of the exponential phase of recovery at 100 mV
After the initial delay, the following time course of recovery can
be fitted by a single exponential function (Fig.
8A). The recovery rate
(reciprocal of the time constant from the exponential fit) shows no
apparent voltage dependence and remains ~3.5
sec1 between 100 and 200 mV (Fig.
8B). The mean recovery rate at 90 mV is also in the
same range, but there is more variability of the data than those at the
other voltages (Fig. 8B, larger error bar at 90
mV). The increased variability at 90 mV may be ascribable to the
small recovered current, which is usually no more than 20% of the
control current in pulse 1 and is always only a few tens of
picoamperes in amplitude. It is therefore hard to conclude
whether the recovery rate at 90 mV (the most positive among the
feasible recovery potentials) is the same as the recovery rates at more
negative potentials or whether it is actually slower than the saturated
recovery rate, like the cases in fibroblasts or in cloned 1G T-type
channels (Chen and Hess 1990; Serrano et al., 1999) (but see Herrington
and Lingle, 1992). In either case, the voltage-independent saturated
recovery rate at 100 mV and more negative potentials is again
qualitatively similar to but quantitatively distinct from the case of
Na+ channels, in which the saturated
recovery rate is achieved only with potentials more negative than 180
mV (Kuo and Bean, 1994). Because additional hyperpolarization can no
longer speed the recovery from inactivation, at saturating negative
membrane potentials the channels seem to recover from a conformation
with the lowest affinity to the inactivating peptide. Along with the
gradually shortened initial delay with increasing hyperpolarization in
Figure 7, these findings are also consistent with the foregoing notion that the horizontal transitions in Scheme 1 (e.g., OB to CB) are voltage-dependent, yet the vertical transitions (e.g., CB to C) lack
intrinsic voltage dependence.
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Figure 8.
The exponential phase of recovery from
inactivation of T-channels over a wide range of membrane potentials.
A, Recovery of the inactivated T-channels is assessed by
a similar two-pulse protocol to that in Figure 7, but here the duration
of recovery gap potential (Vr) is
much longer for study of the later phase of recovery. Treatment of the
data with the aforementioned smoothing technique is no longer necessary
because the fraction of recovered channels here is generally much
larger than that obtained with the very short
Vr in Figure 7. The lines are
monoexponential fits of the form: fraction recovered = 0.47 0.47exp(t/353) (for
Vr = 100 mV, and t
denotes length of Vr in milliseconds; the
horizontal axis), fraction recovered = 1 exp(t/349) (for Vr = 150 mV), and fraction recovered = 1 exp(t/340) (for Vr = 200 mV). B, The recovery rates from inactivation are
given by reciprocals of the time constants from the monoexponential
fits in A. The average results from three to six cells
in each different Vr are shown.
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DISCUSSION |
The deactivation-first route for the recovery of
inactivated T-channels
We have demonstrated that the recovery time course of inactivated
T-channels begins with a delay, which is shortened with increasing
hyperpolarization (down to 200 mV) and is followed by an exponential
phase. Also, there is no discernible Ca2+
current associated with recovery at 80 or 120 mV. These findings indicate that, over a wide range of recovery potentials, T-channels always take the deactivation-first route to recover from inactivation and are therefore more similar to Na+
channels (Kuo and Bean, 1994) than to Shaker
K+ channels (Kuo, 1997). This is
interesting considering that the subunits of T-channels and of
Na+ channels both consist of four domains
(repeats) connected into one single peptide chain (Noda et al., 1986;
Noda and Numa, 1987; Perez-Reyes et al., 1998), whereas a functioning
Shaker K+ channel is comprised of four subunits (four separate peptide chains, each corresponding to one
repeat in the subunit of T-channels or
Na+ channels) (Kamb et al., 1987;
MacKinnon, 1991).
Much slower binding and unbinding rates of the inactivating peptide
in T-channels than in Na+ channels
Despite of the foregoing qualitative similarity, there are
significant quantitative differences between T-channels and
Na+ channels. The macroscopic rate of
development of inactivation in Na+
channels in rat hippocampal neurons saturates at ~3.5
msec1 at +100 mV or more positive
potentials (Kuo and Bean, 1994; Kuo and Liao, 2000), whereas in
thalamic neuronal T-channels, the corresponding rate saturates at
~0.07 msec1 at potentials as negative
as 40 mV. According to Scheme 1, these data indicate a ~50-fold
difference in the O to OB rate (the binding rate of the inactivating
peptide) between these two channels. Moreover, in
Na+ channels the "saturated" time
constant of the exponential phase of recovery is ~4
msec1, which is achieved with very
negative potentials close to 200 mV (Kuo and Bean, 1994). However, in
thalamic neuronal T-channels, such a saturated rate is ~1200-fold
slower (~3.5 sec1) and is obtained
with much more positive potentials (e.g., 100 mV). Based on the
hinged-lid model, these data suggest that both forward and backward
gating movements of the inactivating peptide, namely binding to the
most favorable receptor conformation in the activated channel or
unbinding from the most unfavorable conformation in the deactivated
channel, are much slower in T-channels than in
Na+ channels. This would raise the
possibility that binding and unbinding of the inactivation peptide onto
or from these channels are not free diffusion processes (which may be
the case in the ball-and-chain model and Shaker
K+ channels) (Hoshi et al., 1990; Zagotta
et al., 1990) but are processes controlled by conformational changes of
the channel protein. The kinetic data thus suggest a much more
"rigid" hinge in T-channels. (For convenience, we would discuss our
data in this study with the hinged-lid model. Most discussion will
remain valid and be readily translated into other terms or concepts if one prefers the other model. For example, "much more rigid hinge" would be equivalent to much greater inertia of the coupling mechanism between activation and inactivation based on more general
considerations.)
Relatively small difference between binding affinity of the
inactivating peptide to the activated and that to the deactivated
T-channels
We have demonstrated that the recovery rate of inactivated
T-channels in thalamic neurons saturates at 100 mV. Similar findings were also observed in other preparations, such as fibroblasts, clonal
pituitary cells, and cloned 1G channels (Chen and Hess, 1990;
Herrington and Lingle, 1992; Serrano et al., 1999). Based on Figure
9A, scheme 2, it
seems that most inactivated T-channels are readily redistributed to
state C1B (the fully deactivated-blocked state) at 100 mV. In this
regard, it should be noted that sizable T-current is elicited by a test
pulse at 70 mV (Fig. 1C), indicating significant
distribution of unblocked T-channels to state O at this voltage.
Redistribution of most unblocked T-channels to state C1 therefore would
necessitate potentials more negative than 70 mV. Distribution of most
blocked (inactivated) T-channels to state C1B and distribution of most
unblocked T-channels to state C1 thus does not require very different
potentials. This is in sharp contrast with
Na+ channels (Kuo and Bean, 1994), in
which the recovery rate is saturated at 180 to 200 mV, far more
negative than the voltage range at which
Na+ current starts to be significantly
elicited (approximately 50 to 40 mV). The difference between the
binding affinity of inactivating peptide to the activated channel and
that to the fully deactivated channel thus might be quite smaller in
T-channels than in Na+ channels. This
would indicate a much less drastic conformational change in the
inactivation machinery (receptor, hinge, etc.) during the
activation-deactivation process in T-channels.
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Figure 9.
Gating schemes and comparison of the voltage
dependence of the recovery rates between Na+ and
T-channels. A, scheme 2 is a more
elaborate gating diagram than Scheme 1. Depolarization moves the
channel from the fully deactivated state C1 to the activated state O
through intermediate closed states C2 to Cn. The binding affinity
between the blocking inactivating peptide and its receptor presumably
is larger if the channel is more activated (faster binding rate and
slower unbinding rate toward state O, represented by the size of the
vertical arrows). Channel inactivation is thus coupled
to activation. B, The gradually changing recovery rate
between 80 and 180 mV in Na+ channels (the
dashed line and the dashed vertical axis
on the left) (Kuo and Bean, 1994) is ascribable to rapid
redistribution of the inactivated channels among CnB to C1B during the
hyperpolarizing recovery pulse. More negative potentials shift the
distribution more to the left and thus increase the macroscopic
recovery rate, which is saturated at approximately 200 mV when most
channels are in the fully deactivated state C1B. The dashed
line is a Boltzmann function: recovery rate
(msec1) = 4.5/(1 + exp((V + 140)/16)), where V denotes the membrane potential in
millivolts (the horizontal axis) (Kuo and Bean, 1994).
In contrast, the recovery rate of inactivated T-channels in thalamic
neurons saturates at much less negative potentials (approximately 100
mV; the bold line and the bold vertical
axis on the right; also note the 1000-fold
difference between the units of the left and
right vertical axes). The bold line is a
rough estimate (for the rationales underlying the derivation of the
bold line, please refer to Discussion) and is a
Boltzmann function: recovery rate (sec1) = 3.5/(1 + exp((V + 85)/6)), where V
denotes the membrane potential in millivolts. The bold
line is redrawn with a different scale (thin
line and the thin vertical axis on the
right) to demonstrate that the conformational changes
represented by the bold line may actually be just one
small part of the conformational changes represented by the
dashed line. C, scheme 3
is modified from scheme 2 and may be a more appropriate
gating diagram for T-channels. Note that the unbinding and binding
rates (size of the vertical arrows) are unchanged among
different closed states because the binding affinity of the
inactivating particle toward different closed states C1 to Cn
presumably does not change significantly. This scheme may well explain
many key observations on T-channels in this study, including early
saturation of the exponential recovery phase, apparently steeper slope
of the curve describing the voltage dependence of the recovery rate,
and OB to CnB transition as the rate-limiting step in the overall
deactivation process of the inactivated channel.
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A more abrupt conformational change in the inactivation machinery
associated with opening and closing of the T-channel pore
We have noted significant distribution of unblocked T-channels to
state O at 70 mV. Because of higher affinity of the inactivating peptide to the open state than to the closed state, there should be
more significant or even predominant occupancy of state OB for the
(blocked) inactivated T-channels at 70 mV. This is consistent with
the observation that there is little recovery of T-current at 80 mV
or more positive potentials. Along with the saturated recovery rate at
100 mV, we propose possible voltage-dependence of the recovery rate
of T-channels in Figure 9B (bold line). It is
interesting to note that the curve has a steeper slope than that of the
Na+ channel (dashed line) (Kuo
and Bean, 1994) and that of the inactivation curve in Figure
4C. Because distribution of channels between C1B and OB is
unlikely to involve more charges than distribution between C1 and OB,
the apparently steeper slope would suggest involvement of fewer
intermediate states (in terms of different unbinding rates of the
inactivating peptide) in the distribution of T-channels between C1B and
OB. In other words, T-channels may have significantly less gradual
changes of the inactivation machinery during the activation-deactivation processes than
Na+ channels.
The initial delay is still shortened with increasing hyperpolarization
(Fig. 7A,B), whereas the
exponential phase is saturated. Mechanistically, the delay means the
average time the channel in state OB has to spend before it reaches the
state at which the inactivating peptide could readily unbind. The delay
therefore may serve as an indicator of the deactivation kinetics of
inactivated T-channels. It is interesting that the decaying kinetics of
the tail current, which presumably represents deactivation of the unblocked channels or the O to Cn transition (Fig.
9A), show a similar voltage dependence (0.8 equivalent
charges) (Fig. 5B) to that of the initial delay
(deactivation of the blocked channels, 0.6 equivalent charges) (Fig.
7C). Moreover, the decaying time constant of the tail
T-current is ~4.6 msec at 100 mV. Because of higher affinity of the
inactivating peptide to the O state, OB to CnB rate (Fig.
9A) should be slower. If this is not the rate-limiting step
in the overall transition from OB to C1B, there must be other slower
steps between CnB and C1B. This would be difficult to reconcile with an
initial delay of only ~13.5 msec at 100 mV (Fig. 7B). In
contrast, the deactivating tail current has a time constant of only
~0.12 msec at 100 mV in Na+ channels
(6°C, deactivation of Na+ channels is
too fast to measure at 25°C), yet the initial delay of recovery is
~6 msec in the same condition (Kuo and Bean, 1994). OB to CnB
transition thus is probably the slowest, rate-limiting step in the
entire deactivation process of inactivated T-channels (but not
necessarily so in Na+ channels) and thus
might involve the most significant conformational change of the
inactivation machinery in the activation-deactivation process. If
after the rate-limiting deactivation step the inactivated T-channel
always recover with the same speed (at 100 mV or more negative
potentials) and if there are indeed fewer intermediate deactivated
states of the inactivated T-channel, the conformational change of the
inactivation machinery associated with channel activation-deactivation may not be significant until pore opening or closing in T-channels (Fig. 9C). This relatively "abrupt" change seems to
represent just one part of what happens in
Na+ channels, in terms of both the total
amount of change and the coupling mechanism of inactivation to each
individual molecular step of activation.
Physiological implications
Thalamic relay neurons generate action potentials in two very
different modes, the relay mode and the burst mode (Llinás and
Jahnsen, 1982; McCormick and Feeser, 1990). T-channels have been
implicated to play an important role in the switch between the two
modes (Jahnsen and Llinás, 1984; Coulter et al., 1989; Huguenard
and Prince, 1992). The burst mode prevails at more hyperpolarized basal
membrane potentials that make many T-channels available, so that enough
T-channels could be activated by mild depolarization to further
depolarize the membrane to the threshold of firing bursts of
Na+ spikes. At more depolarized basal
membrane potentials, most T-channels are inactivated and the
spontaneous repetitive bursts of discharge can no longer be sustained.
The neuron then responds more faithfully to external stimuli with
single spike activities (the relay mode). The foregoing slow kinetics
of redistributing T-channels into and out of the inactivated state thus
might put important constraints on the switch between the burst and the
relay modes. It would require a potential change lasting for at least a
few tens of milliseconds, or preferably a few hundred milliseconds, to
make the switch. This is especially so when the membrane is
hyperpolarized and many inactivated T-channels are recovered through
the CB to C pathway. This is probably part of the reason why duration
of the IPSP in thalamic relay neurons, especially the more
long-lasting late hyperpolarization produced by
GABAB response, appears to play an essential role
in timing the spontaneous oscillation (Jahnsen and Llinás, 1984;
Hirsch and Burnod, 1987; Crunelli and Leresche, 1991; Bal et al.,
1995). If T-channels had as fast kinetics as those of
Na+ channels, then even a very transient
perturbation of the membrane potential might lead into modal switch.
The relative slow kinetics of T-channels in thalamic neurons thus might
be viewed as a built-in molecular design that helps to stabilize the
different physiological presentations of the thalamocortical
oscillation system.
| |
FOOTNOTES |
Received Oct. 23, 2000; revised Dec. 27, 2000; accepted Jan. 2, 2001.
This work was supported by National Science Council, Taiwan, Republic
of China Grant NSC-89-2320-B-002-068.
Correspondence should be addressed to Chung-Chin Kuo, Department of
Physiology, National Taiwan University College of Medicine, 1 Jen-Ai
Road, First Section, Taipei 100, Taiwan. E-mail:
cckuo{at}ha.mc.ntu.edu.tw.
| |
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ABSTRACT
INTRODUCTION
