| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Laboratoire de neurobiologie, Ecole Normale Supérieure, 46, rue d'Ulm, 75005 Paris, France
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
= 1 s) and by its sensitivity to external calcium. Steady-state slow inactivation is voltage dependent around the resting membrane potential (the potential of half-inactivation (V0.5) =-70 mV, slope factor = 7.4 mV) and can reduce the calcium current by up to 50%. Near resting potential, the slow inactivation displays a half-time of induction of tens of seconds. The slow inactivation therefore modulates the availability of T-type calcium channels depending upon recent cell history, providing a mechanism to store information in a time scale of seconds.
(Received 12 September 2003;
accepted after revision 12 December 2003;
first published online 19 December 2003)
Corresponding author R. C. Lambert: Laboratoire de neurobiologie, Ecole Normale Supérieure, 46, rue d'Ulm, 75005 Paris, France. Email: rlambert{at}wotan.ens.fr
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
In this paper, we provide evidence that rat CaV3.1 channels expressed in the HEK 293 cell line undergo slow inactivation. We show that the slow inactivation mechanism operates on a time scale of seconds and that up to half of the population of CaV3.1 channels can be driven into slow-inactivated states at the resting potential or during burst firing. The entry into the slow inactivation is characterized by a different apparent voltage dependence from the fast inactivation, reaching its maximal effect at more depolarized potentials than fast inactivation. In addition, slow inactivation is sensitive to the concentration of the divalent charge carrier. These results are compared to the numerous investigations of slow inactivation of sodium and potassium voltage-gated channels.
Since the pore-forming 
1-subunit of cloned T-type channels generates currents similar to native currents, the description and quantification of the slow inactivation process performed here on expressed CaV3.1 channels should help us to understand better the involvement of these channels in neuronal activity. Indeed, slow inactivation may provide a mechanism for this channel population to store information about previous neuronal activity on a time scale of seconds, endowing the neurone with the ability to integrate its own activity for especially long periods.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
HEK 293 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) complemented with 10% fetal bovine serum, 100 U ml-1 penicillin, and 100 µg ml-1 streptomycin. The stably transfected rat Cav3.1 (1G) cell line was kindly provided by Dr. E. Perez-Reyes. The sequence of the rat CaV3.1 transcript used (Lee et al. 1999) is accessible under GenBank accession no. AF027984. Selection was maintained by adding 250 µg ml-1 G-418 to the culture medium. Cells were dissociated using enzymatic digestion with 0.25% trypsin-1 mM EDTA and mechanical trituration. Cells were then split in 35 mm Petri dishes and used during the 1–3 days following the dissociation. All products were purchased from Gibco–Life Technology–Invitrogen.
Transient transfections were also performed using either the calcium-phosphate method or the GeneJammer kit (Stratagene), with a mixture of the CaV3.1 plasmid and the EGFP plasmid to identify the transfected cells. No significant differences were observed between results obtained with stably or transiently transfected cells.
Electrophysiology
Currents were recorded in the whole-cell configuration of the patch-clamp technique using an Axopatch 200A and pCLAMP 8.1 software (Axon Instruments, Union City, CA, USA), filtered at 5 kHz and sampled at 10 kHz. Pipettes were pulled from borosilicate glass capillaries (TW150F-6, WPI, USA) and had a resistance of 1–2 M
when filled with the recording solution. At least 75% of the series resistance (typically < 10 M) and cell capacitance (typically 20 pF) was compensated. Leak currents and capacity transients were subtracted using a P/4 protocol.
To isolate Ca2+ currents, solutions were designed to suppress endogenous K+ and Cl- currents and to fix the Na+ reversal potential close to 0 mV. The pipette solution contained (mM): 10 Hepes, 2.5 CaCl2, 2 MgCl2, 60 methanesulphonic acid, 10 TEA-OH, 70 N-methyl-D-gluconate, 100 NaOH, and 40 BAPTA, 4 Mg-ATP, 15 phosphocreatine and 25 U ml-1 phosphocreatine kinase; pH was adjusted to 7.2 with TEA-OH (solution osmolality was 300 mosmol kg-1). Recording was only started 5 min after rupturing the patch, to allow dialysis of the intracellular medium by the pipette solution.
In most cases, the external solution contained (mM): 10 Hepes, 2 CaCl2, 1 MgCl2, 130 methanesulphonate, 20 TEA-OH, 20 N-methyl-D-glucamine (NMDG) and 120 NaOH, 10 glucose; pH was adjusted to 7.4 with TEA-OH (osmolality was 310 mosmol kg-1).
For concentrations of external calcium higher than 2 mM, NMDG concentration was reduced to maintain solution osmolality. The absence of any endogenous current activated by the various protocols used in the present work was checked on wild-type HEK cells (not shown). All chemicals were purchased from Sigma-Aldrich.
Experiments were carried out at room temperature: 22–24°C.
Protocols
We explain here the principle of the double-pulse protocols. A proportion of the population of the channels was inactivated by the first depolarizing voltage pulse of variable potential and duration. This inactivating pulse was followed by an interpulse interval with specific durations (
T) at hyperpolarized potentials to allow a proportion of channels to recover from inactivation. Finally, a second depolarizing voltage pulse (the test pulse) to -20 mV was applied to test the number of available channels. We refer to the corresponding evoked current as Itest. The amplitude of Itest was not compared to the amplitude of the current evoked during the inactivating pulse, in order to avoid any accumulation of inactivation when applying successive double-pulses or possible run down of the current, which may happen even in the presence of ATP during very long protocols (i.e. recovery from inactivation with 1 min inactivating pulses lasts around 25 min). Instead, each Itest was compared to a specific consecutive control current (Icontrol) evoked at -20 mV after 5 s at the -100 mV holding potential. The ratio Itest/Icontrol was then calculated.
In Fig. 6, the inactivating pulses were applied at different voltages to compensate for the shift of the potential of half-activation (V0.5) of the activation curves that occurs when external calcium is changed (data not shown). The potential of the inactivating pulse, Vip, was -20 mV for 0.5 and 2 mM Ca2+, -10 mV and -5 mV for 10 and 20 mM Ca2+, respectively.
|
For analysis with Clampfit v6.0.3 (Axon Instruments), current traces were numerically low-pass filtered at 1 kHz. All curve fitting was carried out with KyPlot v2.09 (developed by Dr. K. Yoshioka, KyensLab Inc., Tokyo, Japan) using a least-squares routine (Quasi-Newton). The I–V plot was described by a modified Boltzmann function:
|
| (1) |
|
| (2) |
|
| (3) |
1 and 2 the time constants. The curves of recovery from inactivation were described by a bi-exponential function:
|
| (4) |
1 and 2 are the time constants of the fast and slow exponentials and Af is the relative amplitude of the fast component. An additional scaling factor, a, was used when the recovery from inactivation was not complete after 10 s:
|
| (5) |
|
| (6) |
|
|
|
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Using a depolarizing-step protocol over a range of potentials from –70 to +60 mV and a holding potential of -100 mV, families of CaV3.1 currents were recorded from HEK 293 cells; a typical I–V curve is presented in Fig. 1A. As illustrated in the inset, the most noticeable property of the currents generated by this protocol is their very fast decay. Indeed, CaV3.1 channels display the fastest inactivation among all the low voltage-activated calcium channels. With a step depolarization to -10 mV, inactivation occurs with a time constant of 18 ms (n= 9, Fig. 1B). This time constant is not reduced by stronger depolarizations. The involvement of these channels in neuronal activity is strictly dependent upon their steady-state inactivation. Thus, Fig. 1C shows that only a small proportion of the population of the channels is available at -60 mV, a common resting potential in neurones, whereas a hyperpolarization to -100 mV can fully deinactivate the whole population. Moreover, the availability of these channels is highly time-dependent and is determined by the kinetics of their recovery from the inactivation. To examine this property, a double-pulse protocol (see Methods and Fig. 1D) was performed with an inactivating pulse at -20 mV lasting 500 ms, in order to completely inactivate the channels. The proportion of channels recovering from inactivation during periods at -100 mV of various durations (from 20 ms to 10 s) was estimated from the relative amplitude of the current evoked at -20 mV. The description of the recovery kinetics by a single exponential yielded an unsatisfactory fit (dashed line in Fig. 1Dc); a bi-exponential function gave a significantly better fit (continuous line, Fig. 1Dc, P < 0.0001, see Methods). The average values of the time constants are 51 and 199 ms with a relative amplitude of the fast component of 0.36 (n= 10). The presence of two components appeared to be independent of the duration of the inactivating pulse, since in two cells in which inactivation was induced by shorter depolarizations (100 ms), a bi-exponential function with similar time constants was also necessary to fit the data (the time constants for the two cells were, respectively, 25 and 203 ms; and 32 and 235 ms).
Slow inactivation of CaV3.1
Previous investigations in our laboratory have suggested the existence of an additional slower inactivation process that can be induced by long depolarizations in native T-type channels of neurones of nodosus ganglion (Bossu & Feltz, 1986). Other studies have also suggested that T-type channels display slow kinetics of inactivation (Herrington & Lingle, 1992; Klöckner et al. 1999; Frazier et al. 2001; Talavera et al. 2003; and see review in Chen & Hess, 1990; Perez-Reyes, 2003). We therefore investigated the recovery from inactivation induced by long depolarizations of 1 min at -20 mV. As illustrated in Fig. 2B and C, the recovery from inactivation after 1-min-long depolarizations is clearly slower than after prepulses of 500 ms. After inactivation of the channels with long pulses, a hyperpolarization lasting more than 3 s at -100 mV was required to record a peak current of similar amplitude to the control current. The recovery time course was satisfactorily described with the sum of two exponentials. The two time constants obtained from the fit are 110 ms and 1.2 s, with relative amplitude of the fast component of 0.55 (n= 9). Note that the fast time constant of 110 ms is similar to the value of 137 ms estimated when fitting with a single exponential the recovery curve from the fast inactivation induced by 500 ms inactivating pulses (Fig. 1D). We thus concluded that the fast component describes the kinetics of the recovery from the fast-inactivated states and that an additional slow process of recovery, with a time constant around 1 s at -100 mV, appears following long depolarizations. This slow inactivation affects about half of the population of CaV3.1 channels.
|
Because fast inactivation is complete within a few hundred milliseconds, other inactivated states with much slower kinetics do not participate in the definition of the decay of the current evoked by depolarization. Therefore, the entry into other slower inactivation processes cannot be observed directly. Since no molecular approach has succeeded in preventing the development of the fast inactivation in T-type channels, we decided to use an indirect method to measure the onset of slow inactivation. Increasing durations of the inactivating pulse were used to induce progressively slow inactivation of the channels (Fig. 3A). A 350 ms period at the holding potential was subsequently applied to allow most of the channels in the fast-inactivated state to recover. The peak current evoked at -20 mV (Fig. 3B) following this hyperpolarization was thus inversely related to the number of channels accumulated in the slow-inactivated state. Figure 3C shows that for inactivating pulse durations below about 1 s, the ratio Itest/Icontrol was quite stable at 0.92, showing that only around 8% of channels are still in the fast-inactivated state after the 350 ms recovery period. Note that above 1 s, this incompleteness of the recovery from fast inactivation also exists, but becomes negligible when the slow inactivation is induced. For prepulse durations longer than 1 s, the ratio decreases until it plateaus at 0.66 with inactivating prepulses longer than 2 min. Inactivating pulses of more than 4 min were tested and the ratios displayed no further decrease (data not shown). The onset of slow inactivation occurred with a time of half-induction of around 10 s. Although the values of the time constants varied from cell to cell a bi-exponential function was necessary to fit the kinetics of the onset of slow inactivation in each cell (Fig. 3C, P < 0.001, see Methods). Mean values of the time constants were 4.5 and 18.5 s with relative amplitude of each component of 0.08 and 0.21, respectively. More than 30% of Cav3.1 channels enter into the slow-inactivated states when depolarizations last more than 1 min. Note that the discrepancy between the 30% slow inactivation estimated with this protocol and the 50% estimated by the recovery from long inactivation pulses is due to the fact that some slow-inactivated channels recover during the 350 ms interpulse at -100 mV. Since the ratio is never zero even for inactivating pulses even longer than 1 min, we suggest that the slow-inactivated state is not an absorbing state and that there exists an equilibrium between fast- and slow-inactivated channels.
Voltage dependence of the slow inactivation
The voltage dependence of entry into slow inactivation was characterized using a double-pulse protocol with 1 min inactivating pulses to different potentials (see diagrams of the protocols in Fig. 4A). We compared the resulting voltage dependence with that of fast inactivation probed with 500 ms pulses in an analogous protocol. We fitted the steady-state inactivation relations with Boltzmann functions. The V0.5 values were quite similar for fast (500 ms pulse) and slow (1 min pulse) inactivation: -69.0 (n= 4) and -65.7 mV (n= 3), respectively. In contrast, the slope factor for slow inactivation, 7.4 mV, was markedly greater than that for fast inactivation, 4.3 mV. Thus, fast and slow inactivations have different apparent voltage dependences. The principal difference is that slow inactivation is less voltage sensitive or, in other words, is sensitive to voltage over a greater range of potentials. This means that slow inactivation can occur at the resting potentials, at which some silent neurones are likely to remain for several seconds.
Figure 4B shows the recovery from the slow inactivation that had been induced at different potentials and addresses the question of the influence of the inactivating prepulse on the recovery rate. For depolarizations at 20 and -20 mV, the recovery from slow inactivation displayed similar time constants and relatives amplitudes (see legend of Fig. 4 for details). For inactivating pulses to -50 and -70 mV, the recovery from long inactivating pulses is faster. However, the change is due to the shift between the two exponential component rather than any change of the time constants: 150 ms in both cases for the fast component and 0.92 and 1.2 s for the slow component when induced at -50 and -70 mV, respectively. The kinetics of recovery from either fast or slow inactivations are therefore independent of the potential at which the channels enter these inactivated states. However, the relative amplitude of the fast component is increased to 0.80 with inactivating pulses below -20 mV compared to the value of 0.55 estimated for pulses above -20 mV. The dependence of the relative amplitudes of these components upon the potential of the inactivating pulse suggests that the fast-inactivated state is favoured at weakly depolarized potentials. This is in agreement with the differences in apparent voltage dependence of the fast and slow inactivations shown in Fig. 4A. This may indicate a slowing of the entry into the slow-inactivated state at less depolarized potentials.
To investigate the effect of membrane potential on the kinetics of entry into slow inactivation, we employed a double-pulse protocol similar to the one described in Fig. 3 but using inactivating prepluses to -20, -50 and -70 mV (Fig. 4C). The minimal value of the ratios Itest/Icontrol for inactivating pulses at -50 and -20 mV were similar, 0.62 and 0.66, respectively, but reached 0.73 at -70 mV, suggesting that fewer channels had entered into the slow-inactivated states. It is worth noting that with inactivating pulses at -70 mV the minimum of the ratio is higher than with -20 mV prepulse, due to the fact that fast inactivation is not complete at this potential. In addition, there is a shift of the half-maximal induction of slow inactivation towards longer values of the prepulse durations when more hyperpolarized inactivating pulses are applied (Fig. 4C). Accordingly, the corresponding time constants of the bi-exponential fit also increase with the hyperpolarization of the inactivating pulse: 5, 16 and 13 s for the first component, and 19, 35 and 41 s for the second, for -20, -50 and -70 mV, respectively. These results suggest that at hyperpolarized potentials both the rate of slow inactivation and its extent are reduced. Moreover, a near-steady-state is achieved at every potential with 1 min inactivating pulses. This confirms that no shift in the ratios estimated in our previous protocols was introduced because steady state of inactivations had not been reached.
The voltage dependence of recovery from slow inactivation was investigated by applying a double-pulse protocol (see Methods), with varying interpulse potentials ranging from -120 to -50 mV (see diagram of the protocol in Fig. 5A). This protocol is not a direct measurement of the voltage sensitivity of recovery from slow inactivation, because fast inactivation is induced during the 350 ms interpulse at various potentials. However, the plateau observed below -90 mV, when no fast inactivation is induced, indicates that a constant number of channels recover from slow inactivation during a 350 ms hyperpolarization between -20 and -90 mV. This suggests that the recovery from slow inactivation is not intrinsically voltage dependent.
|
Modulation of slow inactivation properties by external calcium concentration
In contrast to HVA calcium channels, T-type channels undergo a fast inactivation that is calcium independent. However, numerous studies of the voltage-gated Na+ (Townsend & Horn, 1997) and K+ channels (Demo & Yellen, 1991; Pardo et al. 1992; Gomez-Lagunas & Armstrong, 1994; Levy & Deutsch., 1996a,b; Kiss & Korn, 1998) have shown modulation of their slow inactivation processes by permeant cations. To address this question for the CaV3.1 channel, we induced slow inactivation in different extracellular concentrations of calcium (0.5, 2, 10 and 20 mM). As illustrated for a typical cell in Fig. 6B, the extent of slow inactivation induced by 1 min inactivating prepulses increased with calcium concentration from 29 ± 2% at 0.5 mM[Ca2+]o to 40 ± 3% at 10 mM (P < 0.005 for 2 and 20 mM compared to 0.5 mM and P < 0.001 for 10 mM compared to 0.5 mM; n= 5) with saturation above 10 mM[Ca2+]o.
Slow inactivation occurs during neuronal activity
We have seen that induction of (Fig. 4A) and recovery from slow inactivation (Fig. 5A) occurs at physiological potentials. To estimate whether slow inactivation can also be induced by sustained neuronal activity, repetitive depolarizations to +20 mV were performed, initially from a -100 mV holding potential. The use of such a hyperpolarized holding potential, although non-physiological, was motivated by the need to isolate the effect of the repetitive brief depolarizations from any inactivation arising before or between depolarization at the holding potential. In this protocol, stimulation had to be maintained at high frequency (200 Hz) in order to observe the entry into slow inactivation (Fig. 7A). We then simulated bursting activity with partial repolarization to -50 mV between brief depolarizations to activate Cav3.1 channels in a more physiological way. In this situation, slow inactivation could be observed at much lower frequency (50 Hz in the example presented in Fig. 7A). However, as already mentioned, since slow inactivation is induced when cells are maintained at -50 mV without any activity (see below and Fig. 4A), the consequence of the repetitive depolarizations on the slow inactivation process is difficult to assess in this latter protocol. In these two protocols, the fits of the entry into slow inactivation are similar to those obtained in Fig. 3C with a sustained inactivating pulse to -20 mV, in agreement with the constant properties of the slow inactivation above -20 mV as shown in Fig. 4.
|
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Fast inactivation of CaV3.1 channels
The present study did not focus upon fast inactivation, which has already been characterized (Serrano et al. 1999; Burgess et al. 2002). Our protocols were therefore not optimally designed to characterize this process. Nevertheless, we observed that the recovery from fast inactivation is best described by the sum of two exponentials in our recording conditions. Although unusual, this is in agreement with the data reported by Monteil et al. (2000) who also found a bi-exponential recovery from fast inactivation with similar time constants in human T-type channel isotypes. Does the slower component reflect the slow inactivation process described here? Since the bi-exponential recovery was observed when channel inactivation was induced with 100 ms inactivating pulses, which induce very little slow inactivation in our hands and since their slow recovery component (= 200 ms) was significantly faster than the recovery from slow inactivation we measure (= 1.2 s), we conclude that fast inactivation has two distinct recovery components that are unrelated to the slow inactivation characterized here.
Slow inactivation of CaV3.1 channels
Previous work on native T-type channels has already described slow kinetics of the recovery from inactivation. This was performed in different preparations with very different recording conditions (Bossu & Feltz, 1986; Herrington & Lingle, 1992; for review see Chen & Hess, 1990; Perez-Reyes, 2003) before the cloning of the T-type channels. The results were interpreted as suggesting the existence of different T-type channels. The subsequent characterization of the three cloned isotypes (Perez-Reyes et al. 1998; Cribbs et al. 1998; Lee et al. 1999) confirmed the existence of channels with different kinetics. However, each of them displays both fast and slow inactivations (Klöckner et al. 1999; Frazier et al. 2001; and the authors' unpublished data).
In this study, we describe the properties of slow inactivation of the CaV3.1 channel. This slow inactivation occurs with a half-time of 10 s during maintained depolarizations at -20 mV (see Fig. 3C). Entry into slow inactivation displays two time constants: a few seconds for the faster one and tens of seconds for the slower one. In contrast, the recovery at -100 mV from slow inactivation induced by long inactivating pulses can be described with a single time constant of 1.1 s. This suggests that although the onset of the slow inactivation appears to be a multistep process, the channels use a common pathway to recover from this inactivation, as we will discuss further below. Moreover, the time to recover from this inactivation is not related to the duration of the previous depolarization (see Fig. 2C). For the slow (C-type) inactivation of voltage-gated potassium channels (Choi et al. 1991), similar observations were made with the Shaker B channels, for which the recovery is also virtually insensitive to the duration of previous stimuli (Toib et al. 1998). In contrast, kinetics of the recovery from the slow inactivation of voltage-gated sodium channels (Rudy, 1978; for review see Goldin, 2003) is related to duration of previous activation (Toib et al. 1998).
Conditions in which the fast and slow inactivations are observable vary, however, from one study to another. When investigating the relationship between deactivation and fast inactivation of CaV3.1 channels, Burgess et al. (2002) reported data of recovery from inactivation after 10 s-long prepulses, a value corresponding to the midpoint of induction of slow inactivation in our conditions. However, the recovery from inactivation appeared to be complete after 1 s at -100 mV (see Fig. 4A in Burgess et al. 2002), which suggests that no component with a slow time constant contributes to this recovery. Talavera et al. (2003) have recently reported fast and a slow time constants in the recovery kinetics of CaV3.1 with values close to our results. However, these kinetics were observed in the recovery from inactivation induced by a short prepulse (200 ms), which would not induce slow inactivation in our recording condition. The differences in the concentration of external ions used may explain these discrepancies of slow inactivation properties and it will be further discussed below.
When depolarized, CaV3.1 channels gradually enter into the slow-inactivated states. Although depolarizations were applied for long durations, slow inactivation was never complete. It reaches a maximum with half of the channels in the slow-inactivated states. The incompleteness of the slow inactivation may reflect a partitioning of the channels during the long prepulse into slow- and fast-inactivated states according to their respective kinetics, and/or the ability of these two inactivation processes to interfere. This latter mechanism occurs in sodium channels in which the slow and fast inactivation gates allosterically interfere, there being competition between the pore conformation for slow inactivation and the docking of the fast inactivating particle in the C-termini of the S6 segments of domain I and IV (Wang et al. 2003; but see also Hilber et al. 2001, 2002). Slow inactivation processes also occur in HVA calcium channels and Shi & Soldatov (2002) have similarly suggested that the slow and fast inactivation gates of the CaV2.1 channel interfere, via a mechanism involving a structural motif at the cytoplasmic end of the S6 segments (for review see Soldatov, 2003). The evidence provided by Staes et al. 2001) and Marksteiner et al. (2001) suggests that the fast inactivation gate in the T-type channel could be localized in the vicinity of the C-termini of the S6 segments and the initial part of the C-terminus of the protein. Similarly to what occurs in sodium and HVA calcium channels, the closure of the fast inactivation gate in the intracellular part of CaV3.1 may allosterically impair the development of the structural conformational changes required to reach slow-inactivated states.
Voltage dependence of slow inactivation
Macroscopic fast inactivation is voltage dependent mainly because it is kinetically linked to the activation processes (Droogmans & Nilius, 1989; Chen & Hess, 1990; Serrano et al. 1999). For instance, this is directly observable in Fig. 1C, in which the decay of the current is relatively slow at more negative potentials where activation is incomplete. Similar characteristics were estimated using both our standard protocol with a 1 s inactivating prepulse (Fig. 1B) and the 500 ms inactivating pulse also used in the present study (Fig. 4A). Another protocol was adapted for the slow inactivation properties, with a 1 min prepulse and 350 ms interpulse (see protocol in the inset of Fig. 4A), and thus the results obtained suggests that this inactivation is also coupled to activation (Fig. 4A). However it requires greater depolarizations to reach its maximal effect than the fast inactivation (see Fig. 4A). Similar characteristics are encountered with the slow inactivation of other voltage-dependent channels, in which the requirement of stronger depolarization was related to structural rearrangements. In sodium channels, the absence of an effect of modification in the S5–S6 linkers on the voltage dependence of slow inactivation suggests that the sensitivity to voltage resides outside the pore (Vilin et al. 2001). Indeed, the specific enhancement of the entry into slow-inactivated states induced by mutations in the voltage sensor of domain IV (Mitrovic et al. 2000) provides evidence that the displacement of the voltage sensor and its associated charge movement plays an important role in the slow inactivation processes. Similarly, in Shaker potassium channels, a model of protein rearrangement has been proposed that postulates a molecular coupling of the voltage sensor to the slow-inactivation gate (Olcese et al. 1997; Loots & Isacoff, 1998, 2000). Whether similar structural rearrangements are involved in slow inactivation of CaV3.1 channels and can explain the more depolarized potentials required to induce this inactivation is still to be determined.
Slow inactivation is impaired for small depolarizations at which its onset kinetics are very slow (tens of seconds). One hypothesis to account for this observation is that the slow-inactivated states are reached from the open state and thus tiny depolarizations induce less slow inactivation, with a slower on-rate because activation is limited and slow at those potentials. As a consequence of the fast decay of CaV3.1 current, this pathway would be limited by the availability of the open state during a long depolarization. However, Serrano et al. (1999) have shown that a residual current can be observed during a sustained depolarization as low as -80 mV due to the activation of at most 2% of the channels. Therefore, channels might reach the slow-inactivated state through the open state during sustained depolarizations. An alternative explanation is that a different pathway may exist to reach the slow-inactivated states from non-conducting states such as the closed or fast-inactivated states. During small depolarizations an intermediate conformation with an incomplete charge movement would be favoured, reducing occupation of these slow-inactivated states.
The recovery from fast inactivation is not considered to be intrinsically voltage dependent (Serrano et al. 1999) and Fig. 5A suggests the same property for slow inactivation. Burgess et al. (2002) provided evidence that the channel must deactivate first (with movement of gating charges) before the recovery from inactivation can occur (see also Satin & Cribbs, 2000; Kuo & Yang, 2001). Similarly, the recovery from slow inactivation is probably under the control of the deactivation pathway. Because we measured only a single time constant of recovery from slow inactivation, the microscopic rate of recovery is probably the limiting step in the whole process and is devoid of an intrinsic voltage dependence. Moreover, we have shown that the potential at which the slow-inactivated states are reached has no effect on the time constant of recovery from long inactivating pulses. In conclusion, we suggest that the recovery from slow inactivation always follows the same pathway to leave the slow-inactivated states whatever the pathway used to reach them.
Modulation of slow inactivation by external Ca2+
When the concentration of external calcium is raised, the maximum degree of slow inactivation is increased by 10% between 0.5 and 20 mM. Since our internal solution contains 40 mM BAPTA, an efficient calcium chelator, an intracellular effect of this cation on the induction of slow inactivation is improbable.
Our results may be explained by a change in the equilibrium between the fast and slow inactivations in favour of the latter. In both voltage-gated sodium and potassium channels, the effect of external cations is the opposite. In voltage-gated sodium channels, an increase of the external sodium concentration shifts the steady-state slow inactivation curve in the depolarized direction and accelerates the recovery from this inactivation (Townsend et al. 1997). Similarly, in potassium channels, ions that bind in or near the external mouth of the pore impede the C-type inactivation conformation change (López-Barneo et al. 1993; Baukrowitz & Yellen, 1996; Kiss & Korn, 1998; Kiss et al. 1999) and speed the recovery from this inactivation (Levy & Deutsch, 1996b).
In CaV3.1, the increase in slow inactivation by external calcium saturates at 10–20 mM. Previous studies on native T-type channels have provided evidence that currents saturate when increasing calcium concentration with a KD between 0.33 and 10 mM (Bossu et al. 1985; Carbone & Lux, 1987; Takahashi et al. 1989; Lux et al. 1990; Herrington & Lingle, 1992). Therefore, the saturation of the pore is expected around 20 mM of external calcium. We may hence hypothesize that the selectivity filter is involved in the slow inactivation processes and that the calcium occupancy of the pore stabilizes the slow inactivation conformation of the channel and slows the kinetics of recovery from these inactivated states. Analogously, Talavera et al. (2003) have shown that the mutations of the amino acids EEDD that form the selectivity filter strongly modified the equilibrium between the slow and fast inactivation states. If the ionic occupancy of the pore affects slow inactivation, varying extracellular concentrations of cations that compete for the pore accessibility may explain the apparent discrepancies in inactivation properties of the CaV3.1 channels reported by different teams using various Ca2+ and Na+ extracellular concentration (Burgess et al. 2002; Talavera et al. 2003).
Modulation of neuronal excitability by slow inactivation of CaV3.1 channel
Although the CaV3.1 channel has a limited ability to sustain currents at high frequencies (>20 Hz), and the channels stop conducting after a few spikes (Kozlov et al. 1999), slow inactivation can occur during high-frequency bursting activity. The frequency of spikes within a burst will directly affect the ability to induce slow inactivation and the duration of a burst will determined the extent of slow inactivation. This provides a memory mechanism able to retain information about burst characteristics for several seconds. Furthermore, we provide evidence that in inactive neurones channels can enter slow-inactivated states at the resting potential. Conversely, recovery from slow inactivation will require either prolonged hyperpolarization or repeated bursts of inhibitory events. Because slow inactivation of up to half of the channels can be observed, it is crucial to include this process in our view of the physiological role of CaV3.1 channels in different kinds of neuronal activities, such as the generation of rhythmic activities, rebound potentials and bistability-mediated activities related to the window-current (for review see Huguenard, 1996; Perez-Reyes, 2003).
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Bossu JL & Feltz A (1986). Inactivation of the low-threshold transient calcium current in rat sensory neurones: evidence for a dual process. J Physiol 376, 341–357.
Bossu JL, Feltz A & Thomann JM (1985). Depolarization elicits two distinct calcium currents in vertebrate sensory neurones. Pflugers Arch 403, 360–368.[CrossRef][Medline]
Burgess DE, Crawford O, Delisle BP & Satin J (2002). Mechanism of inactivation gating of human T-type (low-voltage activated) calcium channels. Biophys J 82, 1894–1906.
Carbone E & Lux HD (1987). Kinetics and selectivity of a low-voltage-activated calcium current in chick and rat sensory neurones. J Physiol 386, 547–570.
Chen CF & Hess P (1990). Mechanism of gating of T-type calcium channels. J General Physiol 96, 603–630.
Choi KL, Aldrich RW & Yellen G (1991). Tetraethylammonium blockade distinguishes two inactivation mechanisms in voltage-activated K+ channels. Proc Natl Acad Sci U S A 88, 5092–5095.
Cribbs LL, Lee JH, Yang J, Satin J, Zhang Y, Daud A, Barclay J, Williamson MP, Fox M, Rees M & Perez-Reyes E (1998). Cloning and characterization of alpha1H from human heart, a member of the T-type Ca2+ channel gene family. Circ Res 83, 103–109.
Demo SD & Yellen G (1991). The inactivation gate of the Shaker K+ channel behaves like an open-channel blocker. Neuron 7, 743–753.[CrossRef][Medline]
Droogmans G & Nilius B (1989). Kinetic properties of the cardiac T-type calcium channel in the guinea-pig. J Physiol 419, 627–650.
Frazier CJ, Serrano JR, George EG, Yu X, Viswanathan A, Perez-Reyes E & Jones SW (2001). Gating kinetics of the alpha1I T-type calcium channel. J General Physiol 118, 457–470.
Goldin AL (2003). Mechanisms of sodium channel inactivation. Curr Opin Neurobiol 13, 284–290.[CrossRef][Medline]
Gomez-Lagunas F & Armstrong CM (1994). The relation between ion permeation and recovery from inactivation of ShakerB K+ channels. Biophys J 67, 1806–1815.
Herrington J & Lingle CJ (1992). Kinetic and pharmacological properties of low voltage-activated Ca2+ current in rat clonal (GH3) pituitary cells. J Neurophysiol 68, 213–232.
Hilber K, Sandtner W, Kudlacek O, Glaaser IW, Weisz E, Kyle JW, French RJ, Fozzard HA, Dudley SC & Todt H (2001). The selectivity filter of the voltage-gated sodium channel is involved in channel activation. J Biol Chem 276, 27831–27839.
Hilber K, Sandtner W, Kudlacek O, Schreiner B, Glaaser I, Schutz W, Fozzard HA, Dudley SC & Todt H (2002). Interaction between fast and ultra-slow inactivation in the voltage-gated sodium channel. Does the inactivation gate stabilize the channel structure? J Biol Chem 277, 37105–37115.
Huguenard JR (1996). Low-threshold calcium currents in central nervous system neurons. Annu Rev Physiol 58, 329–348.[CrossRef][Medline]
Kiss L & Korn SJ (1998). Modulation of C-type inactivation by K+ at the potassium channel selectivity filter. Biophys J 74, 1840–1849.
Kiss L, LoTurco J & Korn SJ (1999). Contribution of the selectivity filter to inactivation in potassium channels. Biophys J 76, 253–263.
Klöckner U, Lee JH, Cribbs LL, Daud A, Hescheler J, Pereverzev A, Perez-Reyes E & Schneider T (1999). Comparison of the Ca2+ currents induced by expression of three cloned alpha1 subunits, alpha1G, alpha1H and alpha1I, of low-voltage-activated T-type Ca2+ channels. Eur J Neurosci 11, 4171–4178.[CrossRef][Medline]
Kozlov AS, McKenna F, Lee JH, Cribbs LL, Perez-Reyes E, Feltz A & Lambert RC (1999). Distinct kinetics of cloned T-type Ca2+ channels lead to differential Ca2+ entry and frequency-dependence during mock action potentials. Eur J Neurosci 11, 4149–4158.[CrossRef][Medline]
Kuo CC & Yang S (2001). Recovery from inactivation of T-type Ca2+ channels in rat thalamic neurons. J Neurosci 21, 1884–1892.
Lee JH, Daud AN, Cribbs LL, Lacerda AE, 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.
Levy DI & Deutsch C (1996a). A voltage-dependent role for K+ in recovery from C-type inactivation. Biophys J 71, 3157–3166.
Levy DI & Deutsch C (1996b). Recovery from C-type inactivation is modulated by extracellular potassium. Biophys J 70, 798–805.
Loots E & Isacoff EY (1998). Protein rearrangements underlying slow inactivation of the Shaker K+ channel. J General Physiol 112, 377–389.
Loots E & Isacoff EY (2000). Molecular coupling of S4 to a K+ channel's slow inactivation gate. J General Physiol 116, 623–636.
López-Barneo J, Hoshi T, Heinemann SH & Aldrich RW (1993). Effects of external cations and mutations in the pore region on C-type inactivation of Shaker potassium channels. Receptors Channels 1, 61–71.[Medline]
Lux HD, 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 430, 159–188.
Marksteiner R, Schurr P, Berjukow S, Margreiter E, Perez-Reyes E & Hering S (2001). Inactivation determinants in segment IIIS6 of Cav1. J Physiol 537, 27–34.
Mitrovic N, George AL Jr & Horn R (2000). Role of domain 4 in sodium channel slow inactivation. J General Physiol 115, 707–718.
Monteil A, Chemin J, Leuranguer V, Altier C, Mennessier G, Bourinet E, Lory P & Nargeot J (2000). Specific properties of T-type calcium channels generated by the human alpha 1I subunit. J Biol Chem 275, 16530–16535.
Motulsky HJ & Ransnas LA (1987). Fitting curves to data using nonlinear regression: a practical and nonmathematical review. FASEB J 1, 365–374.[Abstract]
Olcese R, Latorre R, Toro L, Bezanilla F & Stefani E (1997). Correlation between charge movement and ionic current during slow inactivation in Shaker K+ channels. J General Physiol 110, 579–589.
Pardo LA, Heinemann SH, Terlau H, Ludewig U, Lorra C, Pongs O & Stuhmer W (1992). Extracellular K+ specifically modulates a rat brain K+ channel. Proc Natl Acad Sci U S A 89, 2466–2470.
Perez-Reyes E (2003). Molecular physiology of low-voltage-activated T-type calcium channels. Physiol Rev 83, 117–161.
Perez-Reyes E, Cribbs LL, Daud A, Lacerda AE, Barclay J, Williamson MP, Fox M, Rees M & Lee JH (1998). Molecular characterization of a neuronal low-voltage-activated T-type calcium channel. Nature 391, 896–900.[CrossRef][Medline]
Rudy B (1978). Slow inactivation of the sodium conductance in squid giant axons. Pronase resistance. J Physiol 283, 1–21.
Satin J & Cribbs LL (2000). Identification of a T-type Ca2+ channel isoform in murine atrial myocytes (AT-1 cells). Circ Res 86, 636–642.
Serrano JR, Perez-Reyes E & Jones SW (1999). State-dependent inactivation of the alpha1G T-type calcium channel. J General Physiol 114, 185–201.
Shi C & Soldatov NM (2002). Molecular determinants of voltage-dependent slow inactivation of the Ca2+ channel. J Biol Chem 277, 6813–6821.
Soldatov NM (2003). Ca2+ channel moving tail: link between Ca2+-induced inactivation and Ca2+ signal transduction. Trends Pharmacol Sci 24, 167–171.[CrossRef][Medline]
Staes M, Talavera K, Klugbauer N, Prenen J, Lacinova L, Droogmans G, Hofmann F & Nilius B (2001). The amino side of the C-terminus determines fast inactivation of the T-type calcium channel 1G. J Physiol 530, 35–45.
Takahashi K, Tateishi N, Kaneda M & Akaike N (1989). Comparison of low-threshold Ca2+ currents in the hippocampal CA1 neurons among the newborn, adult and aged rats. Neurosci Lett 103, 29–33.[CrossRef][Medline]
Talavera K, Janssens A, Klugbauer N, Droogmans G & Nilius B (2003). Pore structure influences gating properties of the T-type Ca2+ channel 1G. J General Physiol 121, 529–540.
Toib A, Lyakhov V & Marom S (1998). Interaction between duration of activity and time course of recovery from slow inactivation in mammalian brain Na+ channels. J Neurosci 18, 1893–1903.
Townsend C & Horn R (1997). Effect of alkali metal cations on slow inactivation of cardiac Na+ channels. J General Physiol 110, 23–33.
Vilin YY, Fujimoto E & Ruben PC (2001). A single residue differentiates between human cardiac and skeletal muscle Na+ channel slow inactivation. Biophys J 80, 2221–2230.
Wang SY, Bonner K, Russell C & Wang GK (2003). Tryptophan scanning of D1S6 and D4S6 C-termini in voltage-gated sodium channels. Biophys J 85, 911–920.
|
Acknowledgements |
|---|
This article has been cited by other articles:
|
|
 |
|
  M. A. Diana, Y. Otsu, G. Maton, T. Collin, M. Chat, and S. Dieudonne T-Type and L-Type Ca2+ Conductances Define and Encode the Bimodal Firing Pattern of Vestibulocerebellar Unipolar Brush Cells J. Neurosci., April 4, 2007; 27(14): 3823 - 3838. [Abstract] [Full Text] [PDF]
|
|
|
|
, see detail in 
