Role of extracellular Ca2+ in gating of CaV1.2 channels
1 Department of Pharmacology and Physiology, University of Medicine and Dentistry of New Jersey, New Jersey Medical School, 185 South Orange Avenue, Newark, NJ 07103, USA
Abstract
We examined changes in ionic and gating currents in CaV1.2 channels when extracellular Ca2+ was reduced from 10 mM to 0.1 μM. Saturating gating currents decreased by two-thirds (KD 40 μM) and ionic currents increased 5-fold (KD 0.5 μM) due to increasing Na+ conductance. A biphasic time dependence for the activation of ionic currents was observed at low [Ca2+], which appeared to reflect the rapid activation of channels that were not blocked by Ca2+ and a slower reversal of Ca2+ blockade of the remaining channels. Removal of Ca2+ following inactivation of Ca2+ currents showed that Na+ currents were not affected by Ca2+-dependent inactivation. Ca2+-dependent inactivation also induced a negative shift of the reversal potential for ionic currents suggesting that inactivation alters channel selectivity. Our findings suggest that activation of Ca2+ conductance and Ca2+-dependent inactivation depend on extracellular Ca2+ and are linked to changes in selectivity.
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Introduction
Current interpretations of molecular mechanisms of voltage-gated Ca2+ channels assume independence of permeation and gating. However, several studies suggest otherwise. An effect of permeating ions on mean open time is well established (Nelson et al. 1984; Shuba et al. 1991). Permeating ions appear to affect slower components of activation/deactivation kinetics in some reports (Brown et al. 1983; McDonald et al. 1986) but not in others (Fox et al. 1987; Carbone & Lux, 1987; Carbone et al. 1997). Gating currents are altered at low (< 50 μM) extracellular Ca2+ (Brum et al. 1988; Ferreira et al. 2003), and at high (> 50 μM) intracellular Ca2+ (Isaev et al. 2004). The effects of Ca2+ on gating currents were associated with channel inactivation (Charge 1–Charge 2 interconversion, Brum & Rios, 1987; Shirokov et al. 1992). Inactivation is likely to acquire dependence on voltage through its link to activation. Hence, participation of Ca2+ in inactivation at the level of gating currents might indicate a role of Ca2+ in channel activation.
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Our results indicate that extracellular Ca2+ accelerates voltage-dependent steps in channel gating that are required for Ca2+ permeation but not for Na+ permeation. We also demonstrate that Ca2+-dependent inactivation does not reduce currents carried by alkali metal ions. Taken together, these findings suggest that both activation of Ca2+ conductance and Ca2+-dependent inactivation are linked to voltage-dependent steps in channel gating that affect channel selectivity.
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Methods
CaV1.2 channels (1C, 2a and 2a) were transiently expressed in human embryonic kidney tsA-201 cells as previously described (Chien et al. 1995). The intracellular solution contained (mM): 155 CsCl, 10 Hepes, 10 EGTA, 5 Mg-ATP. Extracellular solutions contained (mM): 150 NaCl, 10 Tris-Cl. CaCl2 was added to achieve the necessary [Ca2+]. [Ca2+] below 40 μM was buffered by 10 mM EGTA and measured with a Ca2+-selective electrode. Solutions were at pH 7.2 and 320 mosmol kg–1.
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Symmetric currents were subtracted by the compensation circuitry of the patch-clamp amplifier (Axopatch 200B, Molecular Devices, Union City, CA, USA). The applicability of this technique has been discussed previously (Shirokov et al. 1998). Control pulses to adjust the circuitry were from –90 to –80 mV at 10 mM Ca2+ at the beginning and at the end of each experiment. We discarded tracings if series resistance and/or membrane capacitance changed by more than 5%. Charge transfer was calculated by integrating the current transient after subtraction of the steady-state component determined as a 3 ms average, 10–15 ms after the beginning of the voltage pulse. Curve fitting was done by a non-linear least-squares routine of SigmaPlot (SPSS Inc., Chicago, IL, USA). Data are presented as mean ± S.E.M.
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Results
Figure 1A shows ionic currents elicited by depolarizations from a holding potential of –90 mV in extracellular solutions with 150 mM Na+ and various [Ca2+]. Activation of currents was mono-exponential at Ca2+ concentrations greater than 7 μM, but biphasic activation was observed at 0.1–1.4 μM [Ca2+] (see Supplemental material, Fig. 1). The fast component was in the range (1–4) x 103 s–1and depended on voltage, but not on [Ca2+] (Fig. 1B). The rate of the fast component was not limited by the speed of the voltage clamp (20 x 103 s–1) and was approximately equal to the rate of activation of Ca2+ current at 10 mM Ca2+. The rate of the slow component was about 10 times less and its magnitude increased with both voltage and [Ca2+] (Fig. 1C). The relative contribution of the fast component increased as [Ca2+] decreased (Fig. 1D) suggesting that this component was due to the openings of channels that were not blocked by Ca2+ before the pulse. It follows that the slow component probably reflects Ca2+ unbinding during the voltage pulse.
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A, ionic currents with 150 mM Na+ and different [Ca2+] (indicated) in the extracellular solution. Voltage pulses were applied from –90 mV holding potential to –60 to +50 mV. Red lines illustrate fits by I = A(1 – exp(–kAt)) + B(1 – exp(–kBt)). Traces at 40 μM and 10 mM Ca2+ were fitted by a single exponential. B, rates of the fast component of a bi-exponential fit to activation of currents at submicromolar Ca2+ (coloured symbols) and rates of a single exponential fit to activation of currents at 10 mM Ca2+ (black symbols). The dashed line shows the rates for 10 mM Ca2+ shifted by –40 mV. C, rates of the slow component of a bi-exponential fit to activation of currents at submicromolar Ca2+ (coloured symbols) and rates of a single exponential fit to activation of currents at 40 μM Ca2+ (dark green symbols). The dashed line is the same as in B. D, relative contributions of the fast component.E, magnitudes of the fit. The magnitudes were normalized to the value at 20 mV and 10 mM Ca2+ in each cell. n = 4–7.
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The effects of Ca2+ on ionic and gating currents are compared in Fig. 2. Because gating currents were recorded without blockage of ionic currents, we evaluated them only at voltages that do not open channels, or at the reversal potential of ionic currents (Fig. 2A). Pulses to the reversal potential moved approximately the maximal gating charge. Lowering of extracellular Ca2+ substantially reduced the ON asymmetric capacitive transient at the reversal potential, but it did not significantly change transients elicited below –90 mV.
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A, changes of gating currents at low Ca2+. Traces b are asymmetric currents recorded at 10 mM Ca2+. Traces c are for 1.35 μM Ca2+. Traces shown by thin lines were elicited at –150 mV, thick lines show traces elicited at the reversal potential for ionic currents (70 mV in b; and 25 mV in c). B, dependencies of the peak amplitude of ionic current (Imax, open symbols) and of the Qmax values (filled symbols) on [Ca2+]. Qmax values were determined as described in Supplemental material, Fig. 2. Before averaging, Imax and Qmax were normalized to their values obtained at 10 mM Ca2+ in the same cell. n = 6–16. The Imax values (except the points at 10 and 0.2 mM Ca2+) were fitted by the function I = I + IKD/(KD + [Ca2+]) with parameters: I = 0.33 ± 0.06, I = 5.51 ± 0.93, KD = 0.49 ± 0.08 μM. The Qmax values were fitted by the function Qmax = 1 – QmaxKD/(KD + [Ca2+]) with parameters: Qmax = 0.68 ± 0.11, KD = 39 ± 14 μM. C, intramembrane charge movements recorded from 0 mV holding potential at 10 mM Ca2+ (b) by pulses ranging from –160 to –10 mV, and at 0.1 μM Ca2+ (c) by pulses ranging from –160 to –40 mV. Thick lines highlight traces at –160 mV. D, charge transfer functions for transients recorded as in C. Charges obtained by integration of the ON transients at different voltages were normalized in each cell to the Qmax value obtained by pulses to 70 mV from –90 mV holding potential at 10 mM Ca2+. n = 5. Parameters of the fits by the Boltzmann function at 10 mM Ca2+ are: Q– = –0.77 ± 0.11, Qmax = 0.97 ± 0.08, V1/2 = –95 ± 7 mV, K = 27 ± 4 mV.
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Removal of Ca2+ is expected to cause a –40 mV shift of intramembrane charge movement distribution due to the surface charge screening effect. However, this shift is too small to explain the reduction of charge transfer observed at low Ca2+ (Supplemental material, Fig. 2). A greater negative shift could have happened if channels were inactivated. However, there was no substantial steady-state inactivation of Na+ currents at –90 mV holding potential (Supplemental material, Fig. 3) and there were no significant charge movements at voltages below –90 mV (Fig. 2A), which would occur if gating charge (Charge 1) were to convert to the intramembrane charge movement in inactivated channels (Charge 2).
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Figure 2B plots maximal ionic currents and maximal charge movements at different [Ca2+]. The apparent KD for blocking Na+ currents was about 0.5 μM. Gating charge increased with Ca2+ at much higher concentrations (40 μM). Importantly, maximal Na+ conductance occurred at Ca2+ concentrations where the saturating intramembrane charge movement was reduced by two-thirds. We refer to the charge movement that is sufficient to open channels for Na+ flux as the Q1/3 charge. The low apparent affinity for the effects of Ca2+ on gating charges indicates that Ca2+ acts on voltage-sensing moieties at a site that is different from the high-affinity blocking site in the selectivity filter.
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A loss of intramembrane charge movement at low Ca2+ could be due to voltage sensors that do not move (i.e. they are removed from the electric field) or move too slowly to be detected. The experiment illustrated in Fig. 2C and D demonstrates that all voltage sensors remained mobile at low Ca2+, thus ruling out the first alternative. Here channels were inactivated by prolonged depolarization, so that all intramembrane charge moves as Charge 2 (Brum & Rios, 1987; Shirokov et al. 1992). We then tested whether or not Charge 2 was reduced at low Ca2+. Asymmetric transients were recorded from the holding potential of 0 mV after a brief interpulse to –60 mV (Fig. 2Ca), which is a ‘watershed’ potential between voltage distributions of Charge 1 and Charge 2. Charge transients (Fig. 2C traces b and c) elicited by pulses below –60 mV were similar at 0.1 and 10 mM Ca2+, indicating that all voltage sensors were mobile at low [Ca2+]. Evidently, the portion of charge movement that was reduced in non-inactivated channels (the Q2/3 charge) was too slow to be recorded at the reversal potential. This possibility suggests that at low [Ca2+] a relatively slow voltage-dependent conformational transition may precede the formation of the Ca2+ conductive state(s), whereas these transitions are not required for Na+ conductance (Fig. 1).
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In separate experiments, shown in Fig. 4 of the Supplemental material, we confirmed for cardiac CaV1.2 channels the finding of Brum et al. (1988) in skeletal muscle CaV1.1 channels, that all charge movements are in Charge 2 modality even at negative holding potentials when extracellular solution has no alkali metals and 15 μM Ca2+.
The experiments illustrated in Figs 3 and 4 tested how inactivation at normal Ca2+ affects monovalent cation conductance. The experimental protocol in Fig. 3 involved exchange of extracellular solution during the application of voltage pulses. First, Na+ current was recorded at 1.4 μM Ca2+ (Fig. 3A trace a). Then, the bathing solution was exchanged with one containing 10 mM Ca2+ and the voltage pulse was applied again (red trace b). Ca2+ was removed from the extracellular solution 300 ms after the start of the voltage pulse. During the solution exchange, ionic current at first decayed to nearly zero due to removal of the charge carrier but then recovered due to the time-dependent unblocking of Na+ conductance. Finally, a third voltage pulse was applied at low Ca2+ to record a control Na+ current again (trace c). The important and somewhat unexpected observation was that, despite inactivation of Ca2+ current, Na+ current fully recovered during the second pulse. Although channels inactivated at normal Ca2+ to 46 ± 6% (n = 5) before the solution exchange, the extent of inactivation at low Ca2+ was only 10 ± 3% at that time. Na+ current recovered to 96 ± 8% after the solution exchange, as if there had been no inactivation at normal Ca2+. The extent of restoration was quantified as the ratio of the peak current after removal of Ca2+ to the average of the values of Na+ currents at the time of the peak in control traces acquired before and after application of 10 mM Ca2+.
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A, restoration of Na+ current after inactivation at normal Ca2+. Traces a and c were elicited at 150 mM Na+ and 1.35 mM Ca2+. The red trace b was recorded starting at 150 mM Na+ and 10 mM Ca2+, and then Ca2+ was reduced to 1.35 μM. The dashed line is trace a scaled to the peak of trace b. B, recovery from inactivation at normal Ca2+ was not accelerated by the absence of Ca2+. Trace a was obtained at 10 mM Ca2+. [Ca2+] was changed during the pulses for trace b at the times indicated. Ca2+ current during the second pulse was not affected. Trace c was obtained at 150 mM Na+ and 1.35 μM Ca2+.
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A, comparison of outward currents obtained after 20 mV pre-pulses of different duration. The dotted line in b was obtained by scaling the inward current at 20 mV to go through the peak current at 100 mV after a 50 ms-long pre-pulse. B, determination of the reversal potential after inactivating pre-pulses of different duration. Tracings are shown for the second step ranging from 20 to 90 mV. The dotted line shows current elicited by a 250 ms long pre-pulse to 20 mV and the second step to 90 mV after addition of 10 μM GdCl3 to the bath. C, instantaneous current–voltage relationships after inactivating pre-pulses of different duration as shown in B. Circles plot current magnitudes measured 2 ms after the step from a 12.5 ms-long pre-pulse to 20 mV; triangles plot current magnitudes measured 2 ms after the step from a 250 ms-long pre-pulse to 20 mV. Before averaging, the values were normalized in each cell to current amplitude at 20 mV. n = 6.
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The results illustrated in Fig. 3B are consistent with the idea that the solution change to low [Ca2+] did not induce a recovery of Ca2+ currents from inactivation. Here, two voltage pulses to 0 mV elicited Ca2+ currents at 10 mM Ca2+ (blue trace a); the two pulses were separated by an interpulse interval (1 s) at –40 mV. Some recovery of Ca2+ current occurred during the interpulse interval but current at the second pulse to 0 mV was still substantially smaller than the first because of persisting inactivation. The same voltage pulses were then applied again, but extracellular Ca2+ was temporarily removed, and then restored, during the interpulse interval (red trace b). Despite Ca2+ removal and re-addition, Ca2+ current during the second pulse was identical to that obtained without the solution change (97 ± 5%, n = 5). Hence, the channels either completely recovered from Ca2+-dependent inactivation during the interpulse and manipulations with extracellular Ca2+ did not cause any additional inactivation, or the channels retained their inactivated state during the solution changes. If the latter is true, then the inactivated channels (Fig. 3A trace b) conducted Na+ equally as well as channels that had not undergone Ca2+-dependent inactivation (traces a and c).
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The experiments shown in Fig. 4 tested whether or not inactivation at high Ca2+ attenuates Ca2+ currents but not alkali metal ion currents. Preferential reduction of inward Ca2+ rather than outward Cs+ currents would result in a shift in reversal potential. To test this prediction, Ca2+ currents evoked by long and short pre-pulses to 20 mV were followed by a pulse to 100 mV (Fig. 4A). If the outward current amplitude were simply proportional to the open probability at the end of the pre-pulse, then the current at 100 mV after the long pre-pulse should decrease in parallel with increasing inactivation of the current and no change in reversal potential would be noted. However, the reversal potential after 250 ms-long pre-pulses (44.7 ± 1 mV) was more negative than after 12.5 ms-long pre-pulses (62.2 ± 1 mV), suggesting that the inward Ca2+ currents were inactivated to a greater degree than the outward Cs+ currents. All outward currents were likely to go through the expressed Ca2+ channels, since there were no significant Cs+ outward currents in non-transfected cells and the contribution of non-specific outward currents recorded after blocking Ca2+ currents by 10 μM Gd3+ (dotted line among traces in Fig. 4Bb) was negligible. These data provide compelling evidence that Ca2+-dependent inactivation leads to a preferential inhibition of Ca2+ conductance compared to monovalent cation conductance, i.e. inactivation induces a change in channel selectivity.
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Discussion
Our findings are analogous in many ways to those with Shaker K+ channels. C-type inactivation in Shaker K+ channels is associated with transitions in the channel pore that attenuate conductance, alter channel selectivity (Starkus et al. 1997, 2000), and induce a negative shift of the intramembrane charge movement (Olcese et al. 1997) analogous to Charge 1–Charge 2 interconversion of Ca2+ channels. Extracellular K+ competes with C-type inactivation by binding to a low affinity site formed by external portions of the pore region (Lopez-Barneo et al. 1993; Baukrowitz & Yellen, 1995). Removal of K+ causes dilatation of the selectivity filter in Shaker and subsequent progression to a defunct condition characterized by zero conductance and a 30% reduction of gating charge (Melishchuk et al. 1998; Loboda & Armstrong, 2001). Our results with Ca2+ channels also demonstrate a change in selectivity during inactivation and show that gating currents of CaV1.2 channels are restricted at low Ca2+. Despite these similarities, there are fundamental differences as well, i.e. gating currents are restricted in Ca2+ channels at low Ca2+ before their Na+ conductance is unblocked.
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The fast component of activation that accompanies unblocking of monovalent ion currents (Fig. 1) had not been observed previously. Perhaps this reflects our use of recombinant channels under relatively simple ionic conditions without organic blockers for other types of channels. Because the slow component diminished at low Ca2+, the simplest interpretation of the data is that the fast component reflects openings of channels unoccupied by Ca2+ before depolarization. The slow component of activation might reflect either Ca2+ exit from channels that opened for the first time, slow opening of blocked channels, or both. However, the rate of the slow component of activation (0.1–0.5) x 103 s–1, is less than the exit rate for Ca2+ ions that re-entered already open channels. The entry rate for external Ca2+ re-blocking inward K+ or Li+ currents, is nearly diffusion limited (109 M–1 s–1) and the exit rate is greater than 103 s–1 (Lansman et al. 1986; Pietrobon et al. 1988; Kuo & Hess, 1993). The exit rate of re-entering Ca2+ ions is reduced by depolarization (Lansman et al. 1986; Kuo & Hess, 1993). In contrast, the slow component of activation was accelerated by voltage (Fig. 1C). Therefore, slow opening of blocked channels is a more likely explanation for the slow component of channel activation at low [Ca2+].
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Our data suggest that activation gating occurs in two sequential and kinetically distinct voltage-dependent steps (Supplemental material, Fig. 5). First, the rapid opening of the voltage-dependent gate allows Na+, but not Ca2+, to permeate the channels. This requires only about a third of the total intramembrane charge to move (Fig. 2). The second slower step is likely to reflect unblocking of Na+ current in a portion of channels that were occupied by Ca2+ before depolarization (Fig. 2). Possibly, the channels at submicromolar [Ca2+] open slowly to pass Ca2+. Both the intramembrane charge movement and the second activation step are accelerated by Ca2+, presumably acting at an extracellular site with a relatively low affinity, possibly at the priming site of Brum et al. (1988).
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The assumption that Na+ ions might permeate the channels that had not undergone the slow conformational change that allows Ca2+ conductance is also consistent with the observation that Na+ ions might permeate the channels that are inactivated to prevent Ca2+ conductance (Figs 3 and 4). Current decay during inactivation at normal Ca2+ apparently involves the second step that is required for Ca2+, but not alkali metal ion, conductance. Interestingly, potentiated L-type Ca2+ channels also preferentially conduct inward Ca2+, but not outward Cs+, currents (Leuranguer et al. 2003).
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Regardless of the above speculations, our data allow us to propose that both activation of Ca2+ conductance and Ca2+-dependent inactivation are sensitive to extracellular Ca2+ and involve alterations in channel selectivity.
Supplemental Material
The online version of this paper can be accessed at: DOI:10.1113/jphysiol.2005.086561
http://jp.physoc.org/cgi/content/full/jphysiol.2005.086561/DCI
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and contains supplemental material consisting of five figures. This material can also be found as part of the full-text HTML version available from http://www.blackwell-synergy.com
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Abstract
We examined changes in ionic and gating currents in CaV1.2 channels when extracellular Ca2+ was reduced from 10 mM to 0.1 μM. Saturating gating currents decreased by two-thirds (KD 40 μM) and ionic currents increased 5-fold (KD 0.5 μM) due to increasing Na+ conductance. A biphasic time dependence for the activation of ionic currents was observed at low [Ca2+], which appeared to reflect the rapid activation of channels that were not blocked by Ca2+ and a slower reversal of Ca2+ blockade of the remaining channels. Removal of Ca2+ following inactivation of Ca2+ currents showed that Na+ currents were not affected by Ca2+-dependent inactivation. Ca2+-dependent inactivation also induced a negative shift of the reversal potential for ionic currents suggesting that inactivation alters channel selectivity. Our findings suggest that activation of Ca2+ conductance and Ca2+-dependent inactivation depend on extracellular Ca2+ and are linked to changes in selectivity.
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Introduction
Current interpretations of molecular mechanisms of voltage-gated Ca2+ channels assume independence of permeation and gating. However, several studies suggest otherwise. An effect of permeating ions on mean open time is well established (Nelson et al. 1984; Shuba et al. 1991). Permeating ions appear to affect slower components of activation/deactivation kinetics in some reports (Brown et al. 1983; McDonald et al. 1986) but not in others (Fox et al. 1987; Carbone & Lux, 1987; Carbone et al. 1997). Gating currents are altered at low (< 50 μM) extracellular Ca2+ (Brum et al. 1988; Ferreira et al. 2003), and at high (> 50 μM) intracellular Ca2+ (Isaev et al. 2004). The effects of Ca2+ on gating currents were associated with channel inactivation (Charge 1–Charge 2 interconversion, Brum & Rios, 1987; Shirokov et al. 1992). Inactivation is likely to acquire dependence on voltage through its link to activation. Hence, participation of Ca2+ in inactivation at the level of gating currents might indicate a role of Ca2+ in channel activation.
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Our results indicate that extracellular Ca2+ accelerates voltage-dependent steps in channel gating that are required for Ca2+ permeation but not for Na+ permeation. We also demonstrate that Ca2+-dependent inactivation does not reduce currents carried by alkali metal ions. Taken together, these findings suggest that both activation of Ca2+ conductance and Ca2+-dependent inactivation are linked to voltage-dependent steps in channel gating that affect channel selectivity.
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Methods
CaV1.2 channels (1C, 2a and 2a) were transiently expressed in human embryonic kidney tsA-201 cells as previously described (Chien et al. 1995). The intracellular solution contained (mM): 155 CsCl, 10 Hepes, 10 EGTA, 5 Mg-ATP. Extracellular solutions contained (mM): 150 NaCl, 10 Tris-Cl. CaCl2 was added to achieve the necessary [Ca2+]. [Ca2+] below 40 μM was buffered by 10 mM EGTA and measured with a Ca2+-selective electrode. Solutions were at pH 7.2 and 320 mosmol kg–1.
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Symmetric currents were subtracted by the compensation circuitry of the patch-clamp amplifier (Axopatch 200B, Molecular Devices, Union City, CA, USA). The applicability of this technique has been discussed previously (Shirokov et al. 1998). Control pulses to adjust the circuitry were from –90 to –80 mV at 10 mM Ca2+ at the beginning and at the end of each experiment. We discarded tracings if series resistance and/or membrane capacitance changed by more than 5%. Charge transfer was calculated by integrating the current transient after subtraction of the steady-state component determined as a 3 ms average, 10–15 ms after the beginning of the voltage pulse. Curve fitting was done by a non-linear least-squares routine of SigmaPlot (SPSS Inc., Chicago, IL, USA). Data are presented as mean ± S.E.M.
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Results
Figure 1A shows ionic currents elicited by depolarizations from a holding potential of –90 mV in extracellular solutions with 150 mM Na+ and various [Ca2+]. Activation of currents was mono-exponential at Ca2+ concentrations greater than 7 μM, but biphasic activation was observed at 0.1–1.4 μM [Ca2+] (see Supplemental material, Fig. 1). The fast component was in the range (1–4) x 103 s–1and depended on voltage, but not on [Ca2+] (Fig. 1B). The rate of the fast component was not limited by the speed of the voltage clamp (20 x 103 s–1) and was approximately equal to the rate of activation of Ca2+ current at 10 mM Ca2+. The rate of the slow component was about 10 times less and its magnitude increased with both voltage and [Ca2+] (Fig. 1C). The relative contribution of the fast component increased as [Ca2+] decreased (Fig. 1D) suggesting that this component was due to the openings of channels that were not blocked by Ca2+ before the pulse. It follows that the slow component probably reflects Ca2+ unbinding during the voltage pulse.
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A, ionic currents with 150 mM Na+ and different [Ca2+] (indicated) in the extracellular solution. Voltage pulses were applied from –90 mV holding potential to –60 to +50 mV. Red lines illustrate fits by I = A(1 – exp(–kAt)) + B(1 – exp(–kBt)). Traces at 40 μM and 10 mM Ca2+ were fitted by a single exponential. B, rates of the fast component of a bi-exponential fit to activation of currents at submicromolar Ca2+ (coloured symbols) and rates of a single exponential fit to activation of currents at 10 mM Ca2+ (black symbols). The dashed line shows the rates for 10 mM Ca2+ shifted by –40 mV. C, rates of the slow component of a bi-exponential fit to activation of currents at submicromolar Ca2+ (coloured symbols) and rates of a single exponential fit to activation of currents at 40 μM Ca2+ (dark green symbols). The dashed line is the same as in B. D, relative contributions of the fast component.E, magnitudes of the fit. The magnitudes were normalized to the value at 20 mV and 10 mM Ca2+ in each cell. n = 4–7.
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The effects of Ca2+ on ionic and gating currents are compared in Fig. 2. Because gating currents were recorded without blockage of ionic currents, we evaluated them only at voltages that do not open channels, or at the reversal potential of ionic currents (Fig. 2A). Pulses to the reversal potential moved approximately the maximal gating charge. Lowering of extracellular Ca2+ substantially reduced the ON asymmetric capacitive transient at the reversal potential, but it did not significantly change transients elicited below –90 mV.
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A, changes of gating currents at low Ca2+. Traces b are asymmetric currents recorded at 10 mM Ca2+. Traces c are for 1.35 μM Ca2+. Traces shown by thin lines were elicited at –150 mV, thick lines show traces elicited at the reversal potential for ionic currents (70 mV in b; and 25 mV in c). B, dependencies of the peak amplitude of ionic current (Imax, open symbols) and of the Qmax values (filled symbols) on [Ca2+]. Qmax values were determined as described in Supplemental material, Fig. 2. Before averaging, Imax and Qmax were normalized to their values obtained at 10 mM Ca2+ in the same cell. n = 6–16. The Imax values (except the points at 10 and 0.2 mM Ca2+) were fitted by the function I = I + IKD/(KD + [Ca2+]) with parameters: I = 0.33 ± 0.06, I = 5.51 ± 0.93, KD = 0.49 ± 0.08 μM. The Qmax values were fitted by the function Qmax = 1 – QmaxKD/(KD + [Ca2+]) with parameters: Qmax = 0.68 ± 0.11, KD = 39 ± 14 μM. C, intramembrane charge movements recorded from 0 mV holding potential at 10 mM Ca2+ (b) by pulses ranging from –160 to –10 mV, and at 0.1 μM Ca2+ (c) by pulses ranging from –160 to –40 mV. Thick lines highlight traces at –160 mV. D, charge transfer functions for transients recorded as in C. Charges obtained by integration of the ON transients at different voltages were normalized in each cell to the Qmax value obtained by pulses to 70 mV from –90 mV holding potential at 10 mM Ca2+. n = 5. Parameters of the fits by the Boltzmann function at 10 mM Ca2+ are: Q– = –0.77 ± 0.11, Qmax = 0.97 ± 0.08, V1/2 = –95 ± 7 mV, K = 27 ± 4 mV.
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Removal of Ca2+ is expected to cause a –40 mV shift of intramembrane charge movement distribution due to the surface charge screening effect. However, this shift is too small to explain the reduction of charge transfer observed at low Ca2+ (Supplemental material, Fig. 2). A greater negative shift could have happened if channels were inactivated. However, there was no substantial steady-state inactivation of Na+ currents at –90 mV holding potential (Supplemental material, Fig. 3) and there were no significant charge movements at voltages below –90 mV (Fig. 2A), which would occur if gating charge (Charge 1) were to convert to the intramembrane charge movement in inactivated channels (Charge 2).
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Figure 2B plots maximal ionic currents and maximal charge movements at different [Ca2+]. The apparent KD for blocking Na+ currents was about 0.5 μM. Gating charge increased with Ca2+ at much higher concentrations (40 μM). Importantly, maximal Na+ conductance occurred at Ca2+ concentrations where the saturating intramembrane charge movement was reduced by two-thirds. We refer to the charge movement that is sufficient to open channels for Na+ flux as the Q1/3 charge. The low apparent affinity for the effects of Ca2+ on gating charges indicates that Ca2+ acts on voltage-sensing moieties at a site that is different from the high-affinity blocking site in the selectivity filter.
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A loss of intramembrane charge movement at low Ca2+ could be due to voltage sensors that do not move (i.e. they are removed from the electric field) or move too slowly to be detected. The experiment illustrated in Fig. 2C and D demonstrates that all voltage sensors remained mobile at low Ca2+, thus ruling out the first alternative. Here channels were inactivated by prolonged depolarization, so that all intramembrane charge moves as Charge 2 (Brum & Rios, 1987; Shirokov et al. 1992). We then tested whether or not Charge 2 was reduced at low Ca2+. Asymmetric transients were recorded from the holding potential of 0 mV after a brief interpulse to –60 mV (Fig. 2Ca), which is a ‘watershed’ potential between voltage distributions of Charge 1 and Charge 2. Charge transients (Fig. 2C traces b and c) elicited by pulses below –60 mV were similar at 0.1 and 10 mM Ca2+, indicating that all voltage sensors were mobile at low [Ca2+]. Evidently, the portion of charge movement that was reduced in non-inactivated channels (the Q2/3 charge) was too slow to be recorded at the reversal potential. This possibility suggests that at low [Ca2+] a relatively slow voltage-dependent conformational transition may precede the formation of the Ca2+ conductive state(s), whereas these transitions are not required for Na+ conductance (Fig. 1).
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In separate experiments, shown in Fig. 4 of the Supplemental material, we confirmed for cardiac CaV1.2 channels the finding of Brum et al. (1988) in skeletal muscle CaV1.1 channels, that all charge movements are in Charge 2 modality even at negative holding potentials when extracellular solution has no alkali metals and 15 μM Ca2+.
The experiments illustrated in Figs 3 and 4 tested how inactivation at normal Ca2+ affects monovalent cation conductance. The experimental protocol in Fig. 3 involved exchange of extracellular solution during the application of voltage pulses. First, Na+ current was recorded at 1.4 μM Ca2+ (Fig. 3A trace a). Then, the bathing solution was exchanged with one containing 10 mM Ca2+ and the voltage pulse was applied again (red trace b). Ca2+ was removed from the extracellular solution 300 ms after the start of the voltage pulse. During the solution exchange, ionic current at first decayed to nearly zero due to removal of the charge carrier but then recovered due to the time-dependent unblocking of Na+ conductance. Finally, a third voltage pulse was applied at low Ca2+ to record a control Na+ current again (trace c). The important and somewhat unexpected observation was that, despite inactivation of Ca2+ current, Na+ current fully recovered during the second pulse. Although channels inactivated at normal Ca2+ to 46 ± 6% (n = 5) before the solution exchange, the extent of inactivation at low Ca2+ was only 10 ± 3% at that time. Na+ current recovered to 96 ± 8% after the solution exchange, as if there had been no inactivation at normal Ca2+. The extent of restoration was quantified as the ratio of the peak current after removal of Ca2+ to the average of the values of Na+ currents at the time of the peak in control traces acquired before and after application of 10 mM Ca2+.
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A, restoration of Na+ current after inactivation at normal Ca2+. Traces a and c were elicited at 150 mM Na+ and 1.35 mM Ca2+. The red trace b was recorded starting at 150 mM Na+ and 10 mM Ca2+, and then Ca2+ was reduced to 1.35 μM. The dashed line is trace a scaled to the peak of trace b. B, recovery from inactivation at normal Ca2+ was not accelerated by the absence of Ca2+. Trace a was obtained at 10 mM Ca2+. [Ca2+] was changed during the pulses for trace b at the times indicated. Ca2+ current during the second pulse was not affected. Trace c was obtained at 150 mM Na+ and 1.35 μM Ca2+.
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A, comparison of outward currents obtained after 20 mV pre-pulses of different duration. The dotted line in b was obtained by scaling the inward current at 20 mV to go through the peak current at 100 mV after a 50 ms-long pre-pulse. B, determination of the reversal potential after inactivating pre-pulses of different duration. Tracings are shown for the second step ranging from 20 to 90 mV. The dotted line shows current elicited by a 250 ms long pre-pulse to 20 mV and the second step to 90 mV after addition of 10 μM GdCl3 to the bath. C, instantaneous current–voltage relationships after inactivating pre-pulses of different duration as shown in B. Circles plot current magnitudes measured 2 ms after the step from a 12.5 ms-long pre-pulse to 20 mV; triangles plot current magnitudes measured 2 ms after the step from a 250 ms-long pre-pulse to 20 mV. Before averaging, the values were normalized in each cell to current amplitude at 20 mV. n = 6.
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The results illustrated in Fig. 3B are consistent with the idea that the solution change to low [Ca2+] did not induce a recovery of Ca2+ currents from inactivation. Here, two voltage pulses to 0 mV elicited Ca2+ currents at 10 mM Ca2+ (blue trace a); the two pulses were separated by an interpulse interval (1 s) at –40 mV. Some recovery of Ca2+ current occurred during the interpulse interval but current at the second pulse to 0 mV was still substantially smaller than the first because of persisting inactivation. The same voltage pulses were then applied again, but extracellular Ca2+ was temporarily removed, and then restored, during the interpulse interval (red trace b). Despite Ca2+ removal and re-addition, Ca2+ current during the second pulse was identical to that obtained without the solution change (97 ± 5%, n = 5). Hence, the channels either completely recovered from Ca2+-dependent inactivation during the interpulse and manipulations with extracellular Ca2+ did not cause any additional inactivation, or the channels retained their inactivated state during the solution changes. If the latter is true, then the inactivated channels (Fig. 3A trace b) conducted Na+ equally as well as channels that had not undergone Ca2+-dependent inactivation (traces a and c).
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The experiments shown in Fig. 4 tested whether or not inactivation at high Ca2+ attenuates Ca2+ currents but not alkali metal ion currents. Preferential reduction of inward Ca2+ rather than outward Cs+ currents would result in a shift in reversal potential. To test this prediction, Ca2+ currents evoked by long and short pre-pulses to 20 mV were followed by a pulse to 100 mV (Fig. 4A). If the outward current amplitude were simply proportional to the open probability at the end of the pre-pulse, then the current at 100 mV after the long pre-pulse should decrease in parallel with increasing inactivation of the current and no change in reversal potential would be noted. However, the reversal potential after 250 ms-long pre-pulses (44.7 ± 1 mV) was more negative than after 12.5 ms-long pre-pulses (62.2 ± 1 mV), suggesting that the inward Ca2+ currents were inactivated to a greater degree than the outward Cs+ currents. All outward currents were likely to go through the expressed Ca2+ channels, since there were no significant Cs+ outward currents in non-transfected cells and the contribution of non-specific outward currents recorded after blocking Ca2+ currents by 10 μM Gd3+ (dotted line among traces in Fig. 4Bb) was negligible. These data provide compelling evidence that Ca2+-dependent inactivation leads to a preferential inhibition of Ca2+ conductance compared to monovalent cation conductance, i.e. inactivation induces a change in channel selectivity.
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Discussion
Our findings are analogous in many ways to those with Shaker K+ channels. C-type inactivation in Shaker K+ channels is associated with transitions in the channel pore that attenuate conductance, alter channel selectivity (Starkus et al. 1997, 2000), and induce a negative shift of the intramembrane charge movement (Olcese et al. 1997) analogous to Charge 1–Charge 2 interconversion of Ca2+ channels. Extracellular K+ competes with C-type inactivation by binding to a low affinity site formed by external portions of the pore region (Lopez-Barneo et al. 1993; Baukrowitz & Yellen, 1995). Removal of K+ causes dilatation of the selectivity filter in Shaker and subsequent progression to a defunct condition characterized by zero conductance and a 30% reduction of gating charge (Melishchuk et al. 1998; Loboda & Armstrong, 2001). Our results with Ca2+ channels also demonstrate a change in selectivity during inactivation and show that gating currents of CaV1.2 channels are restricted at low Ca2+. Despite these similarities, there are fundamental differences as well, i.e. gating currents are restricted in Ca2+ channels at low Ca2+ before their Na+ conductance is unblocked.
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The fast component of activation that accompanies unblocking of monovalent ion currents (Fig. 1) had not been observed previously. Perhaps this reflects our use of recombinant channels under relatively simple ionic conditions without organic blockers for other types of channels. Because the slow component diminished at low Ca2+, the simplest interpretation of the data is that the fast component reflects openings of channels unoccupied by Ca2+ before depolarization. The slow component of activation might reflect either Ca2+ exit from channels that opened for the first time, slow opening of blocked channels, or both. However, the rate of the slow component of activation (0.1–0.5) x 103 s–1, is less than the exit rate for Ca2+ ions that re-entered already open channels. The entry rate for external Ca2+ re-blocking inward K+ or Li+ currents, is nearly diffusion limited (109 M–1 s–1) and the exit rate is greater than 103 s–1 (Lansman et al. 1986; Pietrobon et al. 1988; Kuo & Hess, 1993). The exit rate of re-entering Ca2+ ions is reduced by depolarization (Lansman et al. 1986; Kuo & Hess, 1993). In contrast, the slow component of activation was accelerated by voltage (Fig. 1C). Therefore, slow opening of blocked channels is a more likely explanation for the slow component of channel activation at low [Ca2+].
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Our data suggest that activation gating occurs in two sequential and kinetically distinct voltage-dependent steps (Supplemental material, Fig. 5). First, the rapid opening of the voltage-dependent gate allows Na+, but not Ca2+, to permeate the channels. This requires only about a third of the total intramembrane charge to move (Fig. 2). The second slower step is likely to reflect unblocking of Na+ current in a portion of channels that were occupied by Ca2+ before depolarization (Fig. 2). Possibly, the channels at submicromolar [Ca2+] open slowly to pass Ca2+. Both the intramembrane charge movement and the second activation step are accelerated by Ca2+, presumably acting at an extracellular site with a relatively low affinity, possibly at the priming site of Brum et al. (1988).
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The assumption that Na+ ions might permeate the channels that had not undergone the slow conformational change that allows Ca2+ conductance is also consistent with the observation that Na+ ions might permeate the channels that are inactivated to prevent Ca2+ conductance (Figs 3 and 4). Current decay during inactivation at normal Ca2+ apparently involves the second step that is required for Ca2+, but not alkali metal ion, conductance. Interestingly, potentiated L-type Ca2+ channels also preferentially conduct inward Ca2+, but not outward Cs+, currents (Leuranguer et al. 2003).
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Regardless of the above speculations, our data allow us to propose that both activation of Ca2+ conductance and Ca2+-dependent inactivation are sensitive to extracellular Ca2+ and involve alterations in channel selectivity.
Supplemental Material
The online version of this paper can be accessed at: DOI:10.1113/jphysiol.2005.086561
http://jp.physoc.org/cgi/content/full/jphysiol.2005.086561/DCI
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and contains supplemental material consisting of five figures. This material can also be found as part of the full-text HTML version available from http://www.blackwell-synergy.com
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