Correspondence to: Stephen W. Jones, Department of Physiology and Biophysics, Case Western Reserve University, Cleveland, OH 44106., swj{at}po.cwru.edu (E-mail), Fax: 216-368-3952; (fax)
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Abstract |
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We have examined the kinetics of whole-cell T-current in HEK 293 cells stably expressing the 1G channel, with symmetrical Na+i and Na+o and 2 mM Ca2+o. After brief strong depolarization to activate the channels (2 ms at +60 mV; holding potential -100 mV), currents relaxed exponentially at all voltages. The time constant of the relaxation was exponentially voltage dependent from -120 to -70 mV (e-fold for 31 mV;
= 2.5 ms at -100 mV), but
= 1217 ms from-40 to +60 mV. This suggests a mixture of voltage-dependent deactivation (dominating at very negative voltages) and nearly voltage-independent inactivation. Inactivation measured by test pulses following that protocol was consistent with open-state inactivation. During depolarizations lasting 100300 ms, inactivation was strong but incomplete (~98%). Inactivation was also produced by long, weak depolarizations (
= 220 ms at -80 mV; V1/2 = -82 mV), which could not be explained by voltage-independent inactivation exclusively from the open state. Recovery from inactivation was exponential and fast (
= 85 ms at -100 mV), but weakly voltage dependent. Recovery was similar after 60-ms steps to -20 mV or 600-ms steps to -70 mV, suggesting rapid equilibration of open- and closed-state inactivation. There was little current at -100 mV during recovery from inactivation, consistent with
8% of the channels recovering through the open state. The results are well described by a kinetic model where inactivation is allosterically coupled to the movement of the first three voltage sensors to activate. One consequence of state-dependent inactivation is that
1G channels continue to inactivate after repolarization, primarily from the open state, which leads to cumulative inactivation during repetitive pulses.
Key Words: T-channel, cumulative inactivation, recovery from inactivation
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Introduction |
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Voltage-dependent Ca2+ channels provide a pathway for rapid influx of Ca2+ into cells, which plays a crucial role in both electrical and metabolic signaling. Electrophysiological studies have identified two primary channel types, high voltage-activated (HVA)1 and low voltage-activated (LVA, or T-type) channels (1G, which exhibits the key functional properties of T-channels when expressed in Xenopus oocytes (
T-Channels have been distinguished from HVA channels by a set of biophysical properties, including a more negative voltage range for both activation and inactivation, rapid and nearly complete inactivation, and relatively slow channel closing upon repolarization (deactivation) (
Significant heterogeneity has been observed in the kinetics of T-channel gating, particularly inactivation rates and the voltage dependence of steady state inactivation (1G,
1H, and
1I) known to date (
Cloned T-channels have putative S4 transmembrane regions, suggesting that the mechanism of voltage-dependent activation is essentially the same as in other members of the extended family of K+, Na+, and Ca2+ channels. However, little is known about the mechanism of inactivation in T-channels, or its relationship to the various fast and slow voltage-dependent inactivation processes known for other channels. T-channel inactivation has been described either by models based on
The goal of this study was to characterize the gating of T-channels using whole-cell recording from HEK 293 cells stably expressing the 1G clone, with emphasis on the kinetics of inactivation. In this system, it was possible to characterize T-currents over a wide voltage range, under nearly normal ionic conditions (notably, 2 mM Ca2+ as the charge carrier). We found that
1G channels inactivate primarily from the open state, although inactivation at hyperpolarized voltages involves "partially activated" closed states, and the main pathway for recovery from inactivation bypasses the open state. The currents show strong cumulative inactivation in response to repetitive depolarizations, consistent with continued inactivation from the open state even after repolarization.
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Materials and Methods |
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Cell Culture
Generation of the stable HEK 293 cell line expressing rat 1G (sequence data available from EMBL/GenBank/DDBJ under accession no. AF027984) has been described previously (
Electrophysiology
Currents were recorded using conventional whole-cell patch clamp recording, with an Axopatch 200A amplifier and the Clampex program of pClamp v. 6.0.3 (Axon Instruments). The extracellular solution was 140 mM NaCl, 2 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES, adjusted to pH 7.2 with NaOH. The intracellular solution contained 140 mM NaCl, 11 mM EGTA, 2 mM CaCl2, 4 mM MgATP, 1 mM MgCl2, and 10 mM HEPES, pH 7.2 with NaOH. The pipets filled with intracellular solution had resistances of 24 M. The series resistance in the whole-cell configuration (measured from optimal compensation of capacity transients with the amplifier circuitry) was 5.7 ± 0.3 M
, with cell capacitance of 15.3 ± 0.5 pF (n = 26). Series resistance compensation was nominally 8090%. All experiments were performed at room temperature (~20°C).
The holding potential was -100 mV. Currents were recorded on two channels, with on-line leak subtraction using the P/-4 method on one channel, and raw data during depolarizations on the other, to assess the holding current and cell stability. When this is done, Clampex v. 6 incorrectly sets the current to zero at the end of each leak-subtracted record, so all protocols included a significant period of time at the holding potential at the beginning of each record, and the current during that first holding level was set to zero when the leak-subtracted data were analyzed (using Analyze Adjust Baseline for Epoch A in Clampfit).
Data Analysis
Most data analysis used Clampfit v. 6. Exponential fits to data records used the Simplex or Mixed methods of Clampfit. Other curve fitting was done with the Solver function of Microsoft Excel v. 5 or Excel 97. Unless noted otherwise, values are mean ± SEM. For figures showing averaged data, error bars (±SEM) are shown when larger than the symbols.
Since the currents recorded could be >1 nA, data were examined closely for signs of series resistance error. Clamp speed was assessed by the rise time of tail currents, and steady state accuracy by the effect of partial inactivation on the time course of tail currents. For cells used for kinetic analysis of tail currents (e.g., Figure 3), the 1090% rise time was 0.150.35 ms after 10-kHz analogue filtering. Prepulses that caused ~70% inactivation (using the protocol illustrated in the inset to Figure 11) affected the time constant for deactivation at -100 mV by 15%. Since the measured time constants changed 37% per 10 mV near -100 mV (see Figure 5), this suggests
5 mV error.
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Kinetic models were simulated using SCoP (v. 3.51; Simulation Resources). Simulated data were analyzed further using spreadsheets, or were converted to binary files and analyzed with Clampfit.
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Results |
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General Characteristics of Macroscopic Currents
Currents with the properties expected of T-type calcium currents were recorded from HEK 293 cells stably expressing 1G cDNA. Depolarizations in 10-mV increments from a holding potential of -100 mV elicited transient inward and outward currents (Figure 1 A). Currents showed voltage-dependent macroscopic activation and inactivation, with faster kinetics at more depolarized voltages. At intermediate voltages, the currents "cross over" as typically observed for Na+ currents and T-currents (
The ionic conditions used in this study were essentially normal (see MATERIALS AND METHODS), including 2 mM Ca2+o, except that K+i was replaced by Na+i to minimize currents through any endogenous K+ channels that might be present. HEK 293 cells have occasionally been reported to have endogenous ion channels (1G currents are well isolated in our experimental conditions.
The IV curve, measured as in Figure 1 B, is affected both by gating (activation and inactivation) and by permeation (the voltage dependence of ion flow through an open channel). The protocol of Figure 3 A was used to begin to separate those processes. Channels were first activated by a 2-ms pulse to +60 mV, designed to rapidly activate the channels while minimizing inactivation. This protocol allows us to examine permeation, from the instantaneous IV relation measured immediately after repolarization (Figure 3 B). Assuming that the brief step to +60 mV activates the same number of channels each time (consistent with the constancy of the current recorded during the step to +60 mV), the shape of the instantaneous IV should reflect the voltage dependence of current flow through an open channel. This IV is distinctly nonlinear, suggesting complex interactions among permeant ions in the 1G pore. The reversal potential was +24.4 ± 1.3 mV (n = 8), similar to previous reports for native T-channels (
1G is approximately threefold selective for Na+ over Cs+ (
Division of the IV curve (Figure 1 B) by the instantaneous IV curve (Figure 3 B) was used to evaluate the voltage dependence of activation of 1G channels (Figure 4 A). That ratio (PO,r) should be proportional to the number of channels open at the time of peak current at each voltage. Compared with the usual procedure of measuring tail current amplitudes after depolarizations of fixed duration, this method has the advantage of measuring activation at the maximal value for each voltage. The data at
0 mV were fitted to a single Boltzmann function, with half-maximal activation at -48 mV. The data deviate from that function at positive voltages, in part because the current ratios become discontinuous at the reversal potential, but the measured activation was consistently ~20% greater near +60 mV than near 0 mV. For a rapidly inactivating channel, some channels will inactivate before the point of peak inward current, and the extent of that "hidden" inactivation may vary with voltage. Therefore, the activation curve (Figure 4 A) should be considered an empirical measurement, which may not fully describe the true voltage dependence of the microscopic activation process.
The time course of channel activation was nonexponential. At negative voltages, there was a clear sigmoid delay, which could be approximated by m2h or m3h kinetics (not shown). At positive voltages, the initial time course was not well resolved because of a transient outward current, possibly a gating current, which lasted <1 ms at the T-current reversal potential. The time to peak was voltage dependent, changing approximately fourfold over 100 mV from -30 to +70 mV (Figure 4 B).
Voltage Dependence of Inactivation and Deactivation
Macroscopic inactivation was measured by single exponential fits to the time course of current decay using the protocol of Figure 1 (filled symbols, Figure 5 A). Inactivation was relatively slow at more negative voltages (-60 to -40 mV), but varied little with voltage between -30 and +70 mV. One explanation is that the microscopic inactivation process is voltage independent, as proposed for Na+ channels ( = 2.5 ± 0.2 ms at -100 mV). At more depolarized voltages, the time constants varied little (from
= 11.6 ± 0.6 ms at -40 mV to
= 16.6 ± 1.1 ms at +60 mV), and were comparable to the time constants for macroscopic inactivation. These results are consistent with voltage-dependent channel closing, dominating at extreme negative voltages, but nearly voltage-independent inactivation. There was actually a slight increase in the time constant for inactivation with depolarization (~20% from -20 to +60 mV; Figure 5 A).
The rate of T-channel deactivation reaches a voltage-independent limiting rate at extreme negative voltages in some studies (1G, we examined tail currents at voltages as negative as -150 mV. The time constants showed no detectable deviation from exponential voltage dependence (Figure 5 B).
Inactivation and Recovery at Negative Voltages
Substantial inactivation was observed at voltages as negative as -80 mV (Figure 6 A). Pulses to -120 mV had little effect, implying that there is little resting fast inactivation at our holding potential of -100 mV. At -80 mV, inactivation proceeded with = 223 ± 26 ms, and was 70 ± 5% complete (n = 5). Inactivation was nearly complete at -70 mV (94 ± 5%, with
= 237 ± 39 ms, n = 6).
We examined the time and voltage dependence of recovery from inactivation (Figure 6 B), using the protocol illustrated in Figure 6 C. Recovery from inactivation was complete at -100 and -120 mV. Strikingly, the time course was essentially identical at those voltages (Table 1), suggesting voltage-independent recovery from inactivation at voltages where recovery is complete. Recovery was incomplete, but only slightly slower, at -90 and -80 mV (Table 1).
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Figure 6 suggests that inactivation should reach a steady state by ~1 s. To test that, and to measure the properties of steady state inactivation, voltage steps lasting 1 s were given either directly from -100 mV, or after 60-ms steps to -20 mV to inactivate most of the channels (Figure 7). At steady state, the measured channel availability should depend only on the tested voltage, i.e., the channel should have "forgotten" whether the inactivating pulse to -20 mV had been given. This comparison can only be done in a narrow voltage range, near the midpoint of the steady state inactivation curve, where the amplitudes of inactivation and recovery are both measurable. The two protocols gave almost identical availability curves: V1/2 = -82 ± 2 mV, e-fold for 5.3 ± 0.5 mV, amplitude 1.05 ± 0.02 (inactivation); V1/2 = -83 ± 2 mV, e-fold for 4.8 ± 0.1 mV, amplitude 1.06 ± 0.02 (recovery) (n = 6). When the voltage steps lasted <1 s, the measured V1/2 was more negative for the recovery protocol than for inactivation, demonstrating that steady state had not been reached (data not shown).
The time course of inactivation and recovery showed no clear deviation from exponential kinetics for steps lasting up to ~1 s (Figure 6 A). This is consistent with the existence of a single inactivation process for 1G in that time scale. It is possible that separate slow inactivation processes occur in the second-to-minute time scale, as reported for many voltage-dependent channels, so the "steady state" inactivation curve reported here pertains only to the primary "fast" inactivation process.
Completeness of Inactivation
The inactivation curve could be described well by a single Boltzmann relation, assuming that channels inactivate fully at depolarized voltages (Figure 7). The currents recorded during depolarizations do decay to near zero, but small currents are consistently observed at the end of the pulse (Figure 1 A). This was observed even after depolarizations lasting 120 ms (Figure 8 A). If the inactivated state is fully absorbing, only 0.0003 of the channels should remain open after 120 ms (assuming that the current decays toward zero with = 15 ms), but the peak tail current amplitudes correspond to PO,r ~ 0.02 over a wide voltage range (-60 to +70 mV). The tail currents were small and noisy, so the measured current amplitudes show considerable variability, but residual channel activation was clearly detectable.
The completeness of inactivation was evaluated further using longer (300-ms) depolarizations (Figure 8 B). The averaged record shows a small steady state current at -20 mV, followed by a tail current with a fast component appropriate for channel closing at -100 mV. For the five cells included in that record, from a single exponential fit to the first ~30 ms of the tail current, = 2.7 ± 0.5 ms with amplitude 57 ± 11 pA (with an offset of 10 ± 2 pA, discussed below). The PO,r at the peak of the tail current was 0.0118 ± 0.0004 (n = 4). The PO,r estimated from the current at the end of the step to -20 mV was comparable (0.013 ± 0.002, n = 4). These results suggest that inactivation of
1G is strong but only ~99% complete, at least for depolarizations up to 300 ms.
Another possible source of incomplete inactivation is a "window current" produced by overlap of the steady state activation and inactivation curves. Roughly speaking, that current should be maximal halfway between the midpoint voltages of the two curves (approximately -70 mV for 1G). Tail currents after 600-ms pulses to -70 mV were very small (12 ± 5 pA at -100 mV, n = 6), corresponding to a PO,r of ~0.003, suggesting little steady state activation at -70 mV.
As noted above, single exponential fits to tail currents from the protocol of Figure 8 B yielded an apparently nondeactivating component of 10 ± 2 pA, which corresponds to PO,r = 0.002. (Fits to two exponentials gave a slow component of 12 ± 3 pA, = 52 ± 12 ms, with an offset of 4 ± 1 pA, n = 5.) One possible interpretation is that the slow component is a "resurgent current," reflecting channels recovering from inactivation by passing through the open state (
For comparison, we calculated the resurgent current expected if all of the channels must recover through the open state. We used a three-state scheme: C O
I, assuming that channel closing is irreversible at -100 mV. The inactivation (kI) and recovery (k-I) rates can be estimated from the limiting time constants for inactivation (
= 15 ms) and recovery (
= 100 ms): k-I = 1/100 = 0.01 ms-1, and kI = 1/15 - k-I = 0.057 ms-1. From the tail current time constant at -100 mV (
= 2.5 ms), the channel closing rate kC = 1/2.5 - kI = 0.34 ms-1. From the analytic solution to the general three-state model (
= 117 ms), starting from the initial condition PI = 1. Thus, the observed PO,r of 0.002 is consistent with ~8% of the channels recovering through the open state. Since the slow component of the tail current could be explained in other ways (e.g., a small amount of slow deactivation), this value should be considered an upper limit for the fraction of channels that recover through the open state.
Another argument that inactivation and activation are not strictly coupled is that a C ... C O
I scheme predicts much less complete inactivation than observed. If the rate constants for inactivation and recovery are truly voltage independent with the values estimated above, PI = kI/(kI + k-I) = 0.85 at steady state (at depolarized voltages where the C
O reaction favors the open state). This is additional evidence that recovery from inactivation cannot occur primarily through the open state; i.e., the limiting rate for recovery from inactivation is considerably faster than the rate constant for the O
I reaction.
Closed-state Inactivation
Figure 7 demonstrates that there is considerable inactivation at quite negative voltages, below the range where channel activation is detectable (see Figure 1 B and Figure 4). This observation suggests that channels can inactivate directly from closed states. However, it is possible that open-state inactivation could slowly accumulate even if PO is low, perhaps undetectably low. To examine this quantitatively, we calculated the amount of inactivation expected if channels can inactivate only from the open state. That can be done in a model-independent manner, if we make two assumptions: (a) the microscopic rate constant for inactivation kI (O I) is the reciprocal of the nearly voltage-independent time constant measured at more than -30 mV, and (b) recovery from inactivation can be neglected (i.e., inactivation is absorbing). We do not mean to imply that these assumptions are true, but they allow simple calculation of the amount of inactivation expected to be produced by a voltage protocol, and deviations from the "predicted" inactivation are likely to be informative.
The predicted inactivation was calculated as follows: first, after measuring the instantaneous IV relation for a cell (Figure 3 B), currents are converted to relative PO values (PO,r), by dividing the observed current by the instantaneous current at the same voltage. This gives PO,r as a function of time (relative to that at +60 mV). The expected open-state inactivation is then calculated by integrating dPI/dt = kI PO,r . That is calculated as the point-by-point sum
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during the protocol. Note that this calculation does not make any assumptions about the kinetic scheme for channel activation; i.e., it is independent of number and arrangement of closed states. Similar analyses have been done by
At -70 mV, where channel opening was clearly detectable, the observed inactivation was approximately twice the predicted value (Figure 9 A). The difference was larger at -80 mV (Figure 9 B), where inward currents were visible in one or two of the four cells analyzed. If recovery from inactivation were considered, the predicted inactivation would be reduced further, increasing the discrepancy. We conclude that there is excess inactivation that cannot be accounted for by inactivation from the open state, presumably indicating inactivation directly from closed states.
To determine whether inactivation from closed states is a fundamentally different kinetic process from open-state inactivation, we examined recovery from inactivation after 600-ms steps to -70 mV (Figure 10). Recovery from inactivation was similar, whether inactivation was produced primarily from open or closed states (Table 1). Notably, there was little voltage dependence to recovery (varying approximately twofold from -120 to -80 mV), and recovery could be quite rapid ( ~ 100 ms at -120 mV). These results suggest that the inactivated states reached from open and closed states interconvert rapidly. Alternatively, it is possible that a single inactivated state is accessed from both open and closed states.
Open-state Inactivation
The results described above demonstrate that inactivation can occur from closed states, at least for long, weak depolarizations to voltages near the midpoint of the inactivation curve. But what about brief, strong depolarizations? Figure 11 compares the measured and predicted open-state inactivation produced by the protocol of Figure 3. (Inactivation was measured from an additional test pulse to +60 mV, given 20 ms after each record; see Figure 11, inset.) At negative potentials, there is a good match between measured and predicted inactivation. At positive potentials, the predicted inactivation is larger, possibly due to the observed tendency of inactivation to slow slightly at positive voltages, or to some amount of recovery from inactivation.
Most of the inactivation observed at -120 to -100 mV in Figure 11 can be attributed to the predicted open-state inactivation produced during the initial 2-ms step to +60 mV (0.15 ± 0.04, n = 5). But the amount of inactivation increases with depolarization from -90 to -60 mV, and that extra inactivation can be quantitatively explained by inactivation from the open state during the tail current. That is, a fraction of channels inactivate after repolarization, rather than closing. This behavior is expected from inactivation that is state but not voltage dependent, as channels have a "choice" of pathways for leaving the open state (C ... C O
I). In contrast, with a model where inactivation and recovery are intrinsically voltage dependent, channels would begin to recover from inactivation immediately upon repolarization.
Cumulative Inactivation
State-dependent inactivation is often associated with cumulative inactivation, a phenomenon where repetitive pulses produce significant inactivation, even when little or no inactivation is visible during each depolarization (1G, either using square voltage steps (Figure 12 A) or action potentiallike depolarizations (Figure 12 B).
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Cumulative inactivation results from "hidden" inactivation, which can occur either "on the way up" (during activation, before the point of peak current), or "on the way down" (during the tail current). Inactivation "on the way up" is favored if inactivation occurs primarily from intermediate closed states (
Another sign of state-dependent inactivation is "nonmonotonic recovery from inactivation" (
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Kinetic Model
A model for channel gating can be useful both as an empirical description and as a testable hypothesis for the underlying mechanism. We wanted to develop a model that could reproduce the major experimental results of this study: inactivation is state dependent, fastest from open states, but detectable from closed states. Deactivation is strongly voltage dependent, compared with channel opening (measured as time to peak). Inactivation is strong, but there is a sustained current, corresponding to a PO,r of 12%, over a wide voltage range. Inactivation and recovery reach voltage-independent limiting rates. Repetitive depolarizations produce cumulative inactivation, but inactivation is stronger during a single maintained depolarization.
We considered a model where inactivation is coupled allosterically to voltage sensor activation (Figure 14 A), which has proven successful for describing inactivation for several voltage-dependent channels (
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We began by assuming that all four voltage sensors are allosterically coupled to inactivation. Simulations initially appeared to be successful, but close examination revealed an interesting discrepancy. At voltages near the threshold for significant activation, the sustained current was larger than observed experimentally. Although the true steady state PO did increase monotonically with depolarization, after depolarizations producing partial inactivation (e.g., 60120 ms), the simulated tail currents were approximately twice as large at near-threshold negative voltages than at positive voltages. This was not seen experimentally (Figure 8 A; even more clear for 60-ms depolarizations, where the tail currents were larger and more easily measurable; data not shown). In fact, although the characteristic "crossover" (
The crossover exhibited by the simulations suggested that the model underestimated the rate of inactivation at more negative voltages, which presumably must occur from closed states. We thus modified the scheme, so that activation of only the first three voltage sensors affects the inactivation rate. That is, the last voltage sensor to move (C3C4) has no further effect on the inactivation rate, and the open channel inactivates at the same rate as closed channels in C3 and C4. This is arbitrary, but there is precedent for differential coupling of voltage sensors to inactivation of Na+ channels (
The scheme of Figure 14 A can accurately describe many aspects of the experimental data (Figure 15). Current records cross over at negative voltages (Figure 15 A) and activate in the appropriate voltage range (Figure 15B and Figure C). The sum of two Boltzmann distributions was required for accurate description of the simulated activation curve (Figure 15 C; compare with Figure 4 A). The voltage dependence of the time to peak (Figure 15 D) resembled the experimental data (Figure 4 B), approaching 1 ms at strongly positive voltages. Tail currents from the protocol of Figure 3 A decayed nearly monoexponentially (Figure 15E and Figure F), although the model does not describe the small increase in time constant at positive voltages (Figure 5 A). The model reproduces cumulative inactivation (Figure 15 G), with considerable inactivation occurring during tail currents. Nonmonotonic recovery from inactivation occurs after brief (5-ms) steps, although this is barely visible in the P3/P1 ratio (Figure 15 H).
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It is noteworthy that the tail currents could be described by single exponentials (Figure 15E and Figure F), even in the intermediate voltage range (near -60 mV) where some channels inactivate and others deactivate. If both processes were effectively irreversible, a single exponential would result (1/ = kC + kI; see
The model also produced appropriate steady state inactivation, including its steep voltage dependence (V1/2 = 281 mV, e-fold for 5.0 mV). Inactivation near the V1/2 was predominantly from closed states. Recovery from inactivation was weakly voltage dependent (Table 1). There was no obvious resurgent current during recovery from inactivation, but the tail current (primarily reflecting deactivation of the small steady state current) was ~20% slower than after brief depolarizations, reflecting some channels recovering from inactivation through the open state (simulations not shown).
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Discussion |
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Functional expression of the 1G clone in HEK 293 cells produced currents with the essential kinetic properties of T-type calcium currents. Specifically, the voltage dependence of activation (V1/2 ~ -50 mV) is clearly in the LVA range, and inactivation (V1/2 ~ -80 mV) also occurs at more negative voltages than for most HVA channels. Inactivation is not only rapid (
~ 15 ms at -40 mV and above), but also nearly complete.
1G deactivates ~10-fold slower than HVA channels (
= 2.5 ms at -100 mV). Similar properties have been observed for
1G expressed in Xenopus oocytes (
1G currents over a wide voltage range, without detectable contamination from other currents, as shown by the envelope test (Figure 2) and the absence of ionic currents at the observed reversal potential (Figure 3 A).
State Dependence of Inactivation
The main goal of this study was to characterize the kinetics of inactivation in 1G channels. We conclude that inactivation is state dependent, with little intrinsic voltage dependence. For brief strong depolarizations, inactivation occurs primarily from the open state, but long weak depolarizations produce inactivation from partially activated closed states. We will next discuss the evidence for these conclusions.
The macroscopic inactivation and recovery processes reach essentially voltage-independent time constants at extreme voltages, above -50 mV for inactivation and below -90 mV for recovery (Figure 5 and Figure 7). This can be described by intrinsically voltage-dependent inactivation, if rate constants depend nonexponentially on voltage, as for ßh in the original
Although open-state inactivation can account for the effects of brief depolarization (Figure 11), inactivation also occurred slowly during depolarizations to -90 mV (Figure 7), where no channel opening was detectable. At -70 or -80 mV, the amount of observed inactivation considerably exceeded that predicted by voltage-independent open-state inactivation (Figure 9). Unless the rate for open-state inactivation increases more than twofold at these hyperpolarized voltages, which is unlikely, inactivation must also occur from closed states. The simplest explanation for the inactivation observed below -60 mV is that activation of voltage sensors favors inactivation, even if the channel does not open (Figure 14 A). For 1G, inactivation is faster from the open state than from some of the intermediate closed states, since macroscopic inactivation slowed below -40 mV, and a maintained depolarization produced more inactivation than repetitive pulses (Figure 13 A).
Open- and closed-state inactivation of 1G appear to be closely linked processes, since recovery from inactivation is similar after procedures that favor open-state inactivation (60-ms pulses to -20 mV) or closed-state inactivation (600-ms pulses to -70 mV). The absence of a significant inward current during recovery from inactivation (Figure 8) demonstrates that the primary pathway for recovery from inactivation is via closed states.
It is possible that what we describe as open-state inactivation actually occurs from a closed state that is in rapid, voltage-independent equilibrium with the open state ( O equilibrium is strongly to the right for the parameters used, so occupancy of C3 and C4 is generally low.
Although our model describes well many qualitative and quantitative features of the experimental data, it should be considered preliminary. The model parameters were found by trial and error, rather than rigorous parameter estimation procedures based on quantitative error minimization. We have not systematically tested alternative models. Our data do not include information from single-channel or gating current experiments, which have proven important for modeling gating of other channels. We believe it is useful to present this model at this time, as a possible basis for future studies on the gating of both cloned and native T-channels.
Comparison with Native T-currents
1G is likely to underlie native T-currents in some but not all cells. Notably, it is highly expressed in the thalamus (
1 subunits (
1H and
1I) produce T-currents in expression systems (
1 genes cannot be excluded. Other sources of diversity in channel properties, including accessory subunits and posttranslational modifications, remain to be fully explored for T-channels. It has been suggested that
1 subunits normally associated with HVA channels can produce T-like activity under some conditions (
Kinetic and pharmacological diversity among T-channels is well established (
Our data for 1G are consistent with single exponential kinetics both for development of inactivation and for recovery, on a time scale up to 1 s. This is consistent with most previous work on native T-channels, although some studies have reported multiexponential kinetics (
1G may also exhibit a second inactivation process, on a longer time scale (
Comparison with Inactivation in Other Voltage-dependent Channels
There are some similarities among inactivation processes for different voltage-dependent channels. Fast inactivation of Na+ channels and N-type inactivation of K+ channels reach a limiting rate at positive voltages. At intermediate voltages, macroscopic inactivation is voltage dependent due to kinetic coupling to the activation process, which is relatively slow at such voltages. Inactivation is strong but not necessarily 100% complete. Inactivation of 1G channels shares these properties.
One striking difference from Na+ channels is that recovery from inactivation shows little voltage dependence for T-current (Table 1;
Inactivation of 1G was strong but incomplete, with 9899% inactivation over a wide voltage range. There is considerable variability in the extent of inactivation of Na+ channels, 7097% in the squid giant axon (
1G (Figure 8). This effect is not clearly associated with a slower macroscopic inactivation rate in squid axon (
1G (Figure 5 A). The decreased inactivation with strong depolarization was voltage dependent in squid axon (
Fast inactivation of Na+ and K+ channels is believed to occur primarily but not exclusively from open states (1G. This contrasts with slower inactivation processes of some K+ (
Possible Physiological Implications
One of the clearest functional roles of native T-channels is generation of the low threshold spike that underlies bursts of action potentials in (e.g.) thalamic relay neurons (1G currents are fully consistent with such a scheme.
1G exhibited a sustained current, with 12% of the channels remaining open at all voltages above -70 mV (Figure 8). Our kinetic model accounts for that current with a finite, voltage-independent rate of recovery from inactivation. This differs from the "window current" predicted from an overlap between the activation and inactivation curves, which has a bell-shaped PO versus voltage relation (if inactivation is complete at positive voltages, as often assumed), peaking near the foot of the activation curve (
It is not possible to extrapolate directly from results in an expression system to the situation in vivo, but several kinetic properties of 1G could have important physiological consequences. Activation is quite rapid at positive voltages, so any
1G channels not already activated in a low threshold spike might be activated significantly by a single Na+-dependent action potential. After repolarization, slow deactivation will keep the channels open for a few milliseconds, producing maintained Ca2+ entry (as noted by
1G during action potentiallike depolarizations (Figure 13).
The cumulative inactivation critically depends on the state dependence of inactivation, combined with the characteristic slow deactivation of T-channels. Previous models for thalamic T-currents resemble the original
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Acknowledgements |
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We thank Drs. Leanne L. Cribbs and Toni Schneider for preparation of the 1G cell line, and Dr. Christopher J. Lingle for helpful comments on a draft of this paper.
This work was supported in part by National Institutes of Health grant NS24471 to S.W. Jones and HL58728 to E. Perez-Reyes, and by a Howard Hughes Medical Institute grant to Case Western Reserve University School of Medicine.
Submitted: March 3, 1999; Revised: May 19, 1999; Accepted: May 28, 1999.
1used in this paper: HVA, high voltage-activated; LVA, low voltage-activated; kI, rate constant for inactivation from the open state; k-I, rate constant for recovery from inactivation (to the open state); kC, rate constant for channel closing; PO, probability that a channel is open; PO,r, PO relative to that produced by a 2-ms depolarization from -100 to +60 mV; PI, probability that a channel is inactivated
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References |
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---|
Aldrich, R.W. (1981) Inactivation of voltage-gated delayed potassium current in molluscan neurons. A kinetic model. Biophys. J. 36:519-532[Abstract].
Aldrich, R.W., Stevens, C.F. (1983) Inactivation of open and closed sodium channels determined separately. Cold Spring Harbor Symp. Quant. Biol. 48:147-153[Medline].
Armstrong, C.M., Matteson, D.R. (1985) Two distinct populations of calcium channels in a clonal line of pituitary cells. Science. 227:65-67[Medline].
Armstrong, C.M., Bezanilla, F. (1977) Inactivation of the sodium channel. II. Gating current experiments. J. Gen. Physiol. 70:567-590[Abstract].
Bean, B.P. (1981) Sodium channel inactivation in the crayfish giant axon. Must channels open before inactivating? Biophys. J. 35:595-614[Abstract].
Bean, B.P. (1985) Two kinds of calcium channels in canine atrial cells. Differences in kinetics, selectivity, and pharmacology. J. Gen. Physiol. 86:1-30[Abstract].
Bean, B.P. (1989) Classes of calcium channels in vertebrate cells. Annu. Rev. Physiol. 51:367-384[Medline].
Bean, B.P., McDonough, S.I. (1998) Two for T. Neuron. 20:825-838[Medline].
Bezanilla, F., Armstrong, C.M. (1977) Inactivation of the sodium channel. I. Sodium current experiments. J. Gen. Physiol. 70:549-566
Berjukow, S., Doring, F., Froschmayr, M., Grabner, M., Glossmann, H., Hering, S. (1996) Endogenous calcium channels in human embryonic kidney (HEK293) cells. Br. J. Pharmacol. 118:748-754[Abstract].
Bossu, J.-L., Feltz, A. (1986) Inactivation of the low-threshold transient calcium current in rat sensory neurones: evidence for a dual process. J. Physiol. (Camb.). 376:341-357[Abstract].
Cannon, S.C., Corey, D.P. (1993) Loss of Na+ channel inactivation by anemone toxin (ATX II) mimics the myotonic state in hyperkalaemic periodic paralysis. J. Physiol. (Camb.). 466:501-520[Abstract].
Carbone, E., Lux, H.D. (1984) A low voltage-activated, fully inactivating Ca channel in vertebrate sensory neurones. Nature. 310:501-502[Medline].
Carbone, E., Lux, H.D. (1987) Single low-voltage-activated calcium channels in chick and rat sensory neurones. J. Physiol. (Camb.). 386:571-601[Abstract].
Catterall, W.A. (1996) Molecular properties of sodium and calcium channels. J. Bioenerg. Biomembr. 28:219-230[Medline].
Chandler, W.K., Meves, H. (1970) Sodium and potassium currents in squid axons perfused with fluoride solutions. J. Physiol. (Camb.). 211:679-705[Medline].
Chen, C., Hess, P. (1990) Mechanism of gating of T-type calcium channels. J. Gen. Physiol. 96:603-630[Abstract].
Cota, G., Armstrong, C.M. (1989) Sodium channel gating in clonal pituitary cells. The inactivation step is not voltage dependent. J. Gen. Physiol. 94:213-232[Abstract].
Cribbs, L.L., Lee, J.-H., Yang, J., Satin, J., Zhang, Y., Daud, A., Barclay, J., Williamson, M.P., Fox, M., Rees, M., Perez-Reyes, E. (1998) Cloning and characterization of 1H from human heart, a member of the T-type Ca2+ channel gene family. Circ. Res. 83:103-109
Dashti, S.R., Serrano, J.R., Cribbs, L.L., Perez-Reyes, E., Jones, S.W. (1999) Selectivity of the 1G T-type Ca2+ channel. Biophys. J. 76:A409. (Abstr.).
DeCoursey, T.E. (1990) State-dependent inactivation of K+ currents in rat type II alveolar epithelial cells. J. Gen. Physiol. 95:617-649[Abstract].
Fox, A.P., Nowycky, M.C., Tsien, R.W. (1987) Kinetic and pharmacological properties distinguishing three types of calcium currents in chick sensory neurones. J. Physiol. (Camb.). 394:149-172[Abstract].
Fukushima, Y., Hagiwara, S. (1985) Currents carried by monovalent cations through calcium channels in mouse neoplastic B lymphocytes. J. Physiol. (Camb.). 358:255-284[Abstract].
Gillespie, J.I., Meves, H. (1980) The time course of sodium inactivation in squid giant axons. J. Physiol. (Camb.). 299:289-307[Abstract].
Gutnick, M.J., Lux, H.D., Swandulla, D., Zucker, H. (1989) Voltage-dependent and calcium-dependent inactivation of calcium channel currents in identified snail neurones. J. Physiol. (Camb.). 412:197-220[Abstract].
Herrington, J., Lingle, C.J. (1992) Kinetic and pharmacological properties of low-voltage-activated Ca2+ current in rat clonal (GH3) pituitary cells. J. Neurophysiol. 68:213-232
Hess, P., Lansman, J.B., Tsien, R.W. (1986) Calcium channel selectivity for divalent and monovalent cations. Voltage and concentration dependence of single channel current in ventricular heart cells. J. Gen. Physiol. 88:293-319[Abstract].
Hodgkin, A.L., Huxley, A.F. (1952a) The components of membrane conductance in the giant axon of Loligo. J. Physiol. (Camb.). 116:473-496.
Hodgkin, A.L., Huxley, A.F. (1952b) A quantitative description of membrane current and its application to conduction and excitation in nerve. J. Physiol. (Camb.). 117:500-544[Medline].
Hoshi, T., Zagotta, W.N., Aldrich, R.W. (1990) Biophysical and molecular mechanisms of Shaker potassium channel inactivation. Science. 250:533-538[Medline].
Huguenard, J.R. (1996) Low-threshold calcium currents in central nervous system neurons. Annu. Rev. Physiol. 58:329-348[Medline].
Huguenard, J.R., Prince, D.A. (1992) A novel T-type current underlies prolonged Ca2+-dependent burst firing in GABAergic neurons of rat thalamic reticular nucleus. J. Neurosci. 12:3804-3817[Abstract].
Huguenard, J.R., McCormick, D.A. (1992) Simulation of the currents involved in rhythmic oscillations in thalamic relay neurons. J. Neurophysiol. 68:1373-1383
Klemic, K.G., Shieh, C.-C., Kirsch, G.E., Jones, S.W. (1998) Inactivation of Kv2.1 potassium channels. Biophys. J. 74:1779-1789
Klemic, K.G., Kirsch, G.E., Jones, S.W. (1999) Two types of slow inactivation of Kv potassium channels. Biophys. J. 76:A191. (Abstr.).
Kuo, C.C., Bean, B.P. (1994) Na+ channels must deactivate to recover from inactivation. Neuron. 12:819-829[Medline].
Lambert, R.C., McKenna, F., Maulet, Y., Talley, E.M., Bayliss, D.A., Cribbs, L.L., Lee, J.-H., Perez-Reyes, E., Feltz, A. (1998) Low-voltage-activated Ca2+ currents are generated by members of the CaVT subunit family (1G/H) in rat primary sensory neurons. J. Neurosci. 18:8605-8613
Lee, J.-H., Daud, A.N., Cribbs, L.L., Lacerda, A.E., Pereverzev, A., Klockner, U., Schneider, T., Perez-Reyes, E. (1999) Cloning and expression of a novel member of the low voltage-activated T-type calcium channel family. J. Neurosci. 19:1912-1921
Lux, H.D., Carbone, E., Zucker, H. (1990) Na+ currents through low-voltage-activated Ca2+ channels of chick sensory neurones: block by external Ca2+ and Mg2+. J. Physiol. (Camb.). 430:159-188[Abstract].
Marom, S., Levitan, I.B. (1994) State-dependent inactivation of the Kv3 potassium channel. Biophys. J. 67:579-589[Abstract].
Martin, R.L., Cribbs, L.L., Perez-Reyes, E., Hanck, D.A. (1998) Development of inactivation of the new T-channel, 1G, expressed in mammalian cells. Soc. Neurosci. Abstr. 24:1823. (Abstr.).
Meir, A., Dolphin, A.C. (1998) Known calcium channel 1 subunits can form low threshold small conductance channels with similarities to native T-type channels. Neuron. 20:341-351[Medline].
Mitrovic, N., George, A.L., Jr., Horn, R. (1998) Independent versus coupled inactivation in sodium channels. Role of the domain 2 S4 segment. J. Gen. Physiol. 111:451-462
Nakashima, Y.M., Todorovic, S.M., Pereverzev, A., Hescheler, J., Schneider, T., Lingle, C.J. (1998) Properties of Ba2+ currents arising from human 1E and
1Eß3 constructs expressed in HEK293 cells: physiology, pharmacology, and comparison to native T-type Ba2+ currents. Neuropharmacology. 37:957-972[Medline].
Narahashi, T., Tsunoo, A., Yoshii, M. (1987) Characterization of two types of calcium channels in mouse neuroblastoma cells. J. Physiol. (Camb.). 383:231-249[Abstract].
Neher, E., Lux, H.D. (1971) Properties of somatic membrane patches of snail neurones under voltage clamp. Pflügers Arch. 322:35-38[Medline].
Nilius, B., Hess, P., Lansman, J.B., Tsien, R.W. (1985) A novel type of cardiac calcium channel with different calcium sensitivity. Nature. 316:443-446[Medline].
Nowycky, M.C., Fox, A.P., Tsien, R.W. (1985) Three types of neuronal calcium channel with different calcium agonist sensitivity. Nature. 316:440-443[Medline].
Patil, P.G., Brody, D.L., Yue, D.T. (1998) Preferential closed-state inactivation of neuronal calcium channels. Neuron. 20:1027-1038[Medline].
Perez-Reyes, E., Cribbs, L.L., Daud, A., Lacerda, A.E., Barclay, J., Williamson, M.P., Fox, M., Rees, M., Lee, J.-H. (1998) Molecular characterization of a neuronal low-voltage-activated T-type calcium channel. Nature. 391:896-900[Medline].
Raman, I.M., Bean, B.P. (1997) Resurgent sodium current and action potential formation in dissociated cerebellar Purkinje neurons. J. Neurosci. 17:4517-4526
Randall, A.D., Tsien, R.W. (1997) Contrasting biophysical and pharmacological properties of T-type and R-type calcium channels. Neuropharmacology. 36:879-893[Medline].
Shuba, Y.M., Teslenko, V.I., Savchenko, A.N., Pogorelaya, N.H. (1991) The effect of permeant ions on single calcium channel activation in mouse neuroblastoma cells: ion-channel interaction. J. Physiol. (Camb.). 443:25-44[Abstract].
Soong, T.W., Stea, A., Hodson, C.D., Dubel, S.J., Vincent, S.R., Snutch, T.P. (1993) Structure and functional expression of a member of the low voltage-activated calcium channel family. Science. 260:1133-1136[Medline].
Todorovic, S.M., Lingle, C.J. (1998) Pharmacological properties of T-type Ca2+ current in adult rat sensory neurons: effects of anticonvulsant and anesthetic agents. J. Neurophysiol. 79:240-252
Vandenberg, C.A., Bezanilla, F. (1991) Single-channel, macroscopic, and gating currents from sodium channels in the squid giant axon. Biophys. J. 60:1499-1510[Abstract].
Wang, X.-J., Rinzel, J., Rogawski, M.A. (1991) A model of the T-type calcium current and the low-threshold spike in thalamic neurons. J. Neurophysiol. 66:839-850
Williams, S.R., Toth, T.I., Turner, J.P., Hughes, S.W., Crunelli, V. (1997) The `window' component of the low threshold Ca2+ current produces input signal amplification and bistability in cat and rat thalamocortical neurones. J. Physiol. (Camb.). 505:689-705[Abstract].
Zhu, G., Zhang, Y., Xu, H., Jiang, C. (1998) Identification of endogenous outward currents in the human embryonic kidney (HEK 293) cell line. J. Neurosci. Methods. 81:73-83[Medline].