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Copyright © 2000 The Biophysical Society. All rights reserved.
Biophysical Journal, Volume 78, Issue 6, 2959-2972, 1 June 2000

doi:10.1016/S0006-3495(00)76835-4

Channels, Receptors, and Transporters

Expression of the α₂δ Subunit Interferes with Prepulse Facilitation in Cardiac L-type Calcium Channels

Daniela Platano^*, 1, Ning Qin^*, #, 2, Francesca Noceti^*, Lutz Birnbaumer^{*, ‡, §, ¶}, Enrico Stefani^*, †, § and Riccardo Olcese^*, ,  

^* Department of Anesthesiology, UCLA School of Medicine, Los Angeles, California 90095-7115
^† Department of Physiology, UCLA School of Medicine, Los Angeles, California 90095-7115
^‡ Department of Biological Chemistry, UCLA School of Medicine, Los Angeles, California 90095-7115
^§ Department of Brain Research, UCLA School of Medicine, Los Angeles, California 90095-7115
^¶ Molecular Biology Institutes, UCLA School of Medicine, Los Angeles, California 90095-7115
^# INFM UdR Ferrara, Dipartimento di Biologia, Università di Ferrara, 441000 Ferrara, Italy

Address reprint requests to Dr. Riccardo Olcese, Dept. of Anesthesiology, UCLA School of Medicine, BH-509A, CHS, Box 957115, Los Angeles, CA 90095-7115. Tel.: 310-794-7808; Fax: 310-825-6649.

¹ Daniela Platano's present address is Dip. Fisiologia Umana e Generale, Universita’ di Bologna, 40127 Bologna, Italy.

² Ning Qin's present address is Analgesics Research, The R. W. Johnson Pharmaceutical Research Institute, Spring House, PA 19477-0776.

Voltage-dependent calcium channels are heteromultimeric membrane proteins that have been ranked in several classes on the basis of their pharmacological and biophysical properties (see Birnbaumer et al). The central element of a functional Ca²⁺ channel is the pore-forming α₁ subunit, which is responsible for most of the electrical and pharmacological properties of the channel. The α₁ subunit is physiologically expressed in combination with regulatory subunits (β, α₂δ, and γ), able to modulate several channel functions. As Ca²⁺ fluxes through calcium channels, it controls a number of processes (e.g., cell excitability, muscle contraction, enzyme activity, and gene expression). Therefore, the modulation of the channels by accessory subunits and second messengers becomes a crucial event for cell function. Some of the roles of accessory subunits in channel modulation have been determined using heterologous systems of expression with controlled experimental conditions (e.g., Xenopus laevis oocytes) (Neely et al,Olcese et al,Felix et al,Gurnett et al).

The dihydropyridine (DHP)-sensitive class of calcium channels (L-type) is regulated by neurotransmitters and drugs, and by strong depolarizations. In fact, single strong depolarizations, as well as trains of depolarizations, can induce a transient increase of the channel open probability (facilitation) that persists after the triggering stimulus has ceased. The voltage-dependent facilitation of L-type calcium channels has been described in neurons (Artalejo et al,Kavalali and Plummer, 1996) in skeletal muscle (Johnson et al) and in cardiac myocytes (Noble and Shimoni, 1981a,Noble and Shimoni, 1981b,Lee, 1987,Fedida et al,Zygmunt and Maylie, 1990,Pietrobon and Hess, 1990). However, the extent and the kinetic properties of the facilitation greatly vary among different tissues and species, and interestingly, Cens et al did not find voltage-dependent facilitation in rodent cardiac cells. The expression of cloned calcium channels, with their accessory subunit in simplified expression systems, sheds light on possible reasons for this variability. When the cloned L-type α_1C subunit is expressed in Xenopus oocytes, its voltage-dependent facilitation appears to be dependent on the type of β subunit coexpressed. Specifically, the facilitation of the L-type channels takes place when the modulatory β₁, β₃, β₄ subunit (Bourinet et al,Cens et al,Cens et al) or the cardiac β_2a subunit (Dai et al) are coexpressed with the pore-forming α_1C subunit. On the contrary, in the presence of the palmitoylated form of the β_2a subunit (neuronal) facilitation is absent (Cens et al,Cens et al,Qin et al). In some cases, trains of fast depolarizations resembling action potentials were successfully used to induce facilitation, stressing a physiological role for this phenomenon (Cloues et al). In this study we show that, besides the involvement of the β subunit, the α₂δ subunit also modulates the long-lasting voltage-dependent facilitation (Costantin et al) of a cardiac L-type calcium channel expressed in Xenopus oocytes. Although the coexpression of the α₂δ subunit results in a loss of current potentiation (Dai et al), the channels seem to behave as if they were constitutively facilitated (Platano et al). Moreover, we show evidence of an increase in charge movement associated with the facilitation of α_1C+β₃, suggesting that a recruitment of silent channels may contribute to the voltage-dependent potentiation. These results have been previously reported in abstract form (Platano et al).

Throughout our study, the amino terminus deletion mutant (ΔN60) of the rabbit cardiac a_1C was expressed in Xenopus oocytes. This mutant has a better expression in oocytes than the full-length clone, yielding to larger Ca²⁺ currents without changes in properties (Wei et al). The α_1C pore-forming subunit was expressed in combination with the Ca²⁺ channel accessory subunits β₃ and α₂δ (Wei et al,Perez-Reyes et al). The β₃ and the rabbit skeletal muscle α₂δ subunits were subcloned into the pAGA2 vector, derived from pGEM-3 (Promega, Madison, WI), containing an alfalfa mosaic virus translational initiation site and a 3′ poly A tail of 92 As to facilitate the expression in Xenopus oocytes (Sanford et al,Wei et al). Briefly, the full-length cDNA encoding β₃ was amplified by PCR from the original clone in pBS using Pfu DNA polymerase (Stratagene, La Jolla, CA) and primers B2.1 (containing an NcoI site) and B2.2 (containing an XbaI site). The PCR product was digested with NcoI and XbaI and subcloned into the pAGA2 vector digested with the same restriction enzymes. The correctness of the constructs was confirmed by DNA sequencing using the dideoxy chain termination method.

To synthesize cRNA, all the constructs were linearized with HindIII, followed by treatment with 2mg/ml proteinase K and 0.5% SDS at 37°C for 30min to remove traces of activity. After two phenol/chloroform extractions and ethanol precipitation, the templates were suspended in DEPC-treated water to a final concentration of 0.5μg/μl. cRNAs were in vitro-synthesized at 37°C for 1–2h in a volume of 25μl containing 40mM Tris-HCl (pH 7.2), 6mM MgCl₂, 10mM dithiothreitol, 0.4mM each of adenosine triphosphate, guanosine triphosphate, cytosine triphosphate and uridine triphosphate, 0.8mM 7-methyl guanosine triphosphate, and 10 U T7 RNA polymerase (Boehringer Mannheim, Indianapolis, IN). The transcription products were then extracted with phenol/chloroform, precipitated twice with ethanol, and suspended in DEPC-treated water to a concentration of 0.4μg/μl. cRNAs for the different subunits (0.2μg/μl) were mixed in a 1:1 ratio and a volume of 50nl was injected per oocyte.

Oocytes were obtained from adult female Xenopus laevis (from Xenopus One, Ltd., Dexter, MI). Frogs were anesthetized by immersion in water containing 0.15–0.17% tricaine methanesulfonate for ∼20min or until full immobility. The ovaries were removed under sterile conditions by surgical abdominal incision and stage V and VI oocytes were selected. The animals were then killed by decapitation. The animal protocol was performed with the approval of the Institutional Animal Care Committee of the University of California, Los Angeles. One day before injection, oocytes were defolliculated by collagenase treatment (type I, 2mg/ml for 40min at room temperature; Sigma, St. Louis, MO). Oocytes were maintained at 19°C in Barth solution supplemented with 50μg/ml gentamycin. Recordings were done 4–12 days after the RNA injection.

The cut-open oocyte voltage clamp technique (Stefani and Bezanilla, 1998) was used to record both ionic and gating currents from oocytes expressing α_1C Ca²⁺ channels in combination with the regulatory β₃ and α₂δ subunits. The composition of the external solution (recording chamber and guard compartments) was 10mM Ba²⁺, 96mM Na⁺, and 10mM HEPES, titrated to pH 7.0 with methanesulfonic acid (MES). The lower chamber in contact with the part of the oocyte permeabilized with 0.1% saponin, contained 110mM potassium glutamate, and 10mM HEPES titrated to pH 7.0 with NaOH. Before recording, all the oocytes were injected with 100–150nl of BAPTA-Na₄ 50mM, titrated to pH 7.0 with MES to prevent activation of endogenous Ca²⁺ and Ba²⁺ activated Cl⁻ channels (Barish, 1983,Neely et al). For gating current measurements, the ionic current was blocked by replacing 10mM Ba²⁺ in the external solution with 2mM Co²⁺ and 0.2mM La³⁺. To remove contaminating nonlinear charge movement related to the oocytes, endogenous Na/K ATPase (Rakovsky, 1993), 0.1mM ouabain was added to all external solutions. Leakage and linear capacity currents were compensated analogically and subtracted on-line using P/-4 subtraction protocol from −90, −120mV holding potential (SHP). Charge movement was detected for depolarizations more positive than −70mV, and no changes were observed using SHPs of either −90mV or −120mV. These results indicate that negative subtracting pulses from −90 or −120mV SHP are adequate to subtract linear components.

Signals were filtered with an eight-pole Bessel filter to one-fifth of the sampling frequency. All the experiments were performed at room temperature (21–23°C).

Data obtained from Q-V relationships were fitted with single Boltzmann distribution of the form Q_max/{1+exp[z₁F(V_(1/2)1−V_m)/RT]}; G-V curves were fitted by a dual Boltzmann distribution of the form G₁/{1+exp[z₁F(V_half1−V_m)/RT]}+G₂/{1+exp[z₂F(V_half2−V_m)/RT]}, where F and R are the Faraday and gas constant, respectively; T is the absolute temperature; V_half1 and V_half2 are the midpoints of activation; z₁ and z₂ are the effective valences; Q_max the maximum charge, and G₁ and G₂ are the amplitudes of the first and the second components of the distribution. All data are reported as mean values±SEM.

Cardiac α_1C calcium channels and the auxiliary calcium channel β₃ subunit were expressed in Xenopus oocytes with and without the α₂δ subunit. The expression of α_1C+β₃ and α_1C+β₃+α₂δ gave rise to large ionic currents having different properties, depending on the channel subunit composition. Representative traces obtained from the combinations α_1C+β₃ (A) and α_1C+β₃+α₂δ (B) are shown in Fig. 1. The currents were elicited by depolarization to −30mV, 0mV, and +30mV from −90mV holding potential (HP). As previously described by other authors (Bangalore et al,Felix et al,Qin et al) the coexpression of the α₂δ subunit increased the peak current, shifted the current-voltage (I-V) curve peak 10mV to more negative potential, and increased the activation and deactivation rates of α_1C+β₃. These modulatory effects were used as a positive control for the good expression of the α₂δ subunit. When the I-V curves of α_1C+β₃ and α_1C+β₃+α₂δ are normalized to the peak inward current (Figure 1C), the magnitudes of the outward currents are markedly different, while the reversal potentials are practically identical. In the presence of the α₂δ subunit, the outward current had a shallower voltage dependence relative to α_1C+β₃.

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Figure 1

Comparison of voltage-dependent properties of α_1C when expressed in combination with β₃ and β₃+α₂δ. Representative current families from either α_1C+β₃ (A) and α_1C+β₃+α₂δ (B) elicited stepping from −50mV to +50mV in 10-mV increments. The holding potential was −90mV. In (C) are shown normalized I-V curves for α_1C+β₃ (●, n=9) and α_1C+β₃+α₂δ (○, n=6).

We studied the facilitation of the ionic current of α_1C using a double-pulse protocol. The current was recorded during a pulse to 10mV before and after the application of a 200-ms prepulse to +80mV (the protocol is shown on the top panel of Fig. 2). Although the α_1C current did not show potentiation after the prepulse, currents recorded from oocytes expressing α_1C+β₃ were facilitated when preceded by a positive prepulse (200ms at +80mV; Figure 2AB). The potentiation observed for this subunit composition lasted several seconds after the prepulse during a repolarization to −90mV. In Fig. 2, panels A and B show the current facilitation of the subunit composition α_1C+β₃ after a 200-ms prepulse to 80mV followed by repolarization of 50ms (A) and 1s (B) to −90mV. To estimate the duration of the facilitation, the averaged peak facilitated currents (recorded during the test pulse at 10mV) were plotted versus time of repolarization to −90mV (interpulse). The facilitation decayed following a double-exponential time course with time constants of 0.44s and 51.38s. The amplitudes of the two components were, respectively, 55% and 45% of the total facilitation (n=8, Figure 2E).

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Figure 2

The α₂δ subunit seems to prevent prepulse potentiation. (A) α_1C+β₃ superimposed current traces evoked by the voltage protocol shown at top. The current during a 10-mV test pulse increases when the test pulse is preceded by a prepulse to 80mV. The potentiation is long-lasting and still present after a 1-s repolarization to −90mV (B); the voltage protocols are shown above the current traces. When α_1C+β₃ is expressed together with α₂δ, there is no significant potentiation of the current (C) and (D). (E) The extent of potentiation of the ionic current at 10mV is plotted versus the time of repolarization between the prepulse and test pulse for the combination α_1C+β₃. The potentiation decayed in a double-exponential manner with time constants of τ_fast=0.44s, Amp_fast=50.1%, and τ_slow=51.38, Amp_slow=40.8% s, n=8. No significant potentiation was measured by coexpressing the α₂δ subunit (F) (n=6).

When α_1C+β₃ was expressed together with the α₂δ subunit, the current potentiation induced by the prepulse was no longer observed, as shown by the representative current traces in Figure 2CD. In this case, the turn-on of the ionic currents was faster and showed a time-dependent decay. For very brief repolarizations (<50ms) the current during the test pulse was slightly reduced after the positive prepulse, possibly due to inactivation. Figure 2F shows the lack of potentiation in α_1C+β₃+α₂δ: the potentiation after a 200-ms prepulse to 80mV is plotted against the time of repolarization at −90mV (D, n=11). Even stronger prepulses (up to 140mV) did not elicit potentiation in the presence of the α₂δ subunits (data not shown).

By comparing the voltage dependence of the activation (G-V curve) in α_1C+β₃ and α_1C+β₃+α₂δ, and the modulatory effect of positive prepulses, we examined the possibility that the addition of the α₂δ subunit already maximized channel opening, leaving no room for the potentiation process to occur. In this study we used 25-ms depolarizing steps, ranging from −80 to 190mV in 10-mV increments, followed by a repolarization to −50mV delivered every 5s. The G-V curves were constructed plotting the peak tail currents at −50mV against the potentials of the depolarizing steps. The voltage steps were delivered in control conditions (i.e., without prepulse) (Figure 3A) or preceded by a 200-ms prepulse to 80mV (Figure 3B). The recovery from potentiation was measured without prepulses 2–3min later (Figure 3C). As described in Fig. 2, the prepulse increased the ionic current during the test pulse in α_1C+β₃: Figure 3D shows the I-V plot for control (●) and potentiated current (○) obtained by measuring the values of the ionic current at the end of the 25-ms test pulse.

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Figure 3

In α_1C+β₃, a positive prepulse shifts the activation curve to more negative potentials. Three families of Ba²⁺ currents were recorded from an oocyte expressing α_1C+β₃. Representative traces are shown for a 25-ms depolarization to the indicated potentials; tails currents are evoked by repolarization to −50mV. (A) Ionic currents recorded in the absence of a preceding prepulse. (B) the depolarizing steps were preceded by a 200-ms prepulse to 80mV, followed by a repolarization (interpulse) of 50ms to −90mV. Both ionic and gating currents increased in amplitude. (C) The same conditions as in (A); the test protocol without prepulse was run 2min after the experiment shown in (B). (D) Current-voltage relationships from the same oocyte. Note that the I-V curve recorded in the absence of prepulse merged with the potentiated one for potentials above 100mV. (E) Averaged normalized G-V curves measured from the tail currents during the repolarization to −50mV (n=9) for Ba²⁺ current recorded in control (●), after a 200-ms prepulse to +80mV (○), and during the recovery (♦). Error bars represent SEM. Data were fitted by dual Boltzmann distributions (see Methods).

The normalized averaged G-V curves (n=9±SEM) show that the overall effect of the prepulse is to shift the voltage dependence of channel activation to more negative potentials (Figure 3E). The data could be fitted in both control (●) and potentiated (○) G-V values with the sum of two Boltzmann distributions with the same half-activation potentials and slope factors, and with different proportion of the relative amplitudes of the two components. The positive prepulse increased the fraction of the more negative components from 1% to 43% of the total conductance, resulting in an overall negative shift of the G-V curve. The G-V curves are shown normalized to their maxima. The operation was required because of the progressive time-dependent rundown of the conductance, which was enhanced by the demanding pulse protocol. However, as assessed by the experiments shown in Fig. 4, in which the channels were challenged by only two test voltages (to prevent the rundown), the limiting conductance of α_1C+β₃ (measured with a pulse to 180mV) was the same with or without prepulse.

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Figure 4

The limiting conductance of α_1C+β₃ does not change with the prepulse. Current records from an oocyte expressing α_1C+β₃, evoked by two test pulses to +10mV (A) and +180mV (B), preceded or not by a prepulse of 200ms to +80mV. Note the lack of further potentiation when the test pulse is +180mV (B). The bar plot in (C) summarizes the degree of facilitation induced by the prepulse as measured during a test pulse to +10 and +180mV (n=6). The oocytes were challenged with a minimum number of pulses to minimize the rundown of the current. The conditions and the pulse protocol are the same as in Fig. 3.

In oocytes expressing α_1C+β₃+α₂δ, no potentiation of the ionic currents was detected after the positive prepulse (Figure 5A–C). The I-V curves of a representative oocyte are shown in Figure 5D (■, Control; □, Prepulse). The normalized G-V curves obtained from oocytes expressing α_1C+β₃+α₂δ revealed a little effect of the prepulse on the voltage dependency of the channel activation. The G-V curve in α_1C+β₃+α₂δ without prepulse (■) was practically identical to the G-V curve obtained for the combination α_1C+β₃ after the positive prepulse (Figure 5E, dashed line). The dotted line corresponds to the fit of G-V curve of α_1C+β₃ without prepulses. We simultaneously fitted all G-V curves for both α_1C+β₃ and α_1C+β₃+α₂δ, with and without the prepulse, to the same effective valences (z₁ and z₂) and half-activation potentials (V_half1 and V_half2) for the two components, but with a different proportion of their amplitude factors. The effective valences z₁ and z₂ were 1.94 for the first, more negative component, and 0.97 for the second, more positive component. The half-activation potentials (V_half1 and V_half2) were 14.4mV and 93.2mV, respectively. As mentioned for α_1C+β₃ injected oocytes, the first component of the G-V (G₁) increased after the prepulse from 1% of the G_max (in control) to 43% (Figure 3E). However, when α₂δ was coexpressed with α_1C+β₃, the proportion of G₁ was already 53% in control conditions and the prepulse had a minor effect on the ratio between the two amplitudes of the fit (G₁=67% with prepulse). These findings suggest that the presence of the α₂δ sets the channels in a conformational state similar to the one reached after prepulse potentiation.

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Figure 5

In α_1C+β₃+α₂δ, a positive prepulse has little effect on the activation curve. Current records from an oocyte expressing α_1C+β₃+α₂δ are shown. Twenty-five-millisecond depolarizations to the indicated potentials were delivered in control condition (without prepulse) (A) after a 200-ms prepulse to +80mV (B) and 2min after the prepulse protocol (C). The time of repolarization between the prepulse and the test pulse was 50ms at −90mV. In (D) is shown the I-V relationship for the same oocyte in control (■), after the prepulse (□), and after the recovery (△). (E) Averaged and normalized GV curves measured from the tail currents during the repolarization to −50mV (n=6) for Ba²⁺ current recorded in control (■) and after a 200-ms prepulse to +80mV (□). The solid lines are the simultaneous fit to the sum of two Boltzmann distributions. In the plot are also reported the G-V fits of α_1C+β₃ without the prepulse (dotted line) and after prepulse (dashed line) as shown in Fig. 3. Error bars are SEM.

The installation of outward ionic current is much faster in α_1C+β₃+α₂δ than in α_1C+β₃. The slow component of the outward current α_1C+β₃ probably reflects the development of the potentiation occurring during the pulse. We used a 200-ms test pulse to 80mV preceded by an identical prepulse to test this possibility. As shown in Figure 6A, the outward current in α_1C+β₃ during a control pulse to 80mV develops with a relatively slow kinetic. When a prepulse to 80mV is applied, the current during the same test pulse to 80mV develops with a much faster kinetic. The two current traces (with and without prepulse) merge together at the end test pulse, indicating that a ∼150-ms pulse is sufficient to induce the maximal potentiation at 80mV. On the contrary, the coexpression of the α₂δ produces a fast developing ionic current which is virtually unmodified by positive prepulse (Figure 6B).

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Figure 6

Differential effect of a 200-ms prepulse to +80mV on the ionic current from α_1C+β₃ and α_1C+β₃+α₂δ. (A) Two superimposed current traces elicited by a double pulse voltage protocol shown in the top panel. In α_1C+β₃, the current elicited by a test pulse to 80mV develops slowly. The activation kinetics of a pulse to 80mV is greatly accelerated by the application of a 200-ms prepulse to the same potential. After the positive prepulse, the current in the test pulse is already potentiated, as shown by the faster kinetics. Instead, the coexpression with α₂δ (B) elicited a fast-rising current during the test pulse at 80mV (without prepulse), and remains practically unmodified after the prepulse.

Voltage-dependent calcium channels respond to changes in the potential across the plasma membrane by changing their conformational state and giving rise to the gating currents. Thus, the amplitude of gating current is proportional to the number of channels able to gate. To have a simultaneous evaluation of the behavior of the gating current and the change in membrane conductance induced by the prepulse, we recorded the currents at the reversal potential using the standard external solution containing 10mM Ba²⁺ (Fig. 7). By pulsing at the reversal potential, it is possible to record the “ON” gating currents at the beginning of the depolarizing pulse and the ionic tail current partially contaminated by the “OFF” gating current at the end of the pulse, during the repolarization. The experimental reversal potential, ranging from +45mV to +55mV, was determined for each experiment. We found that after the delivery of a 200-ms prepulse to 60mV in the oocytes expressing α_1C+β₃, the facilitation of the ionic current (as assessed by the increase in the tail current) was accompanied by an increase of the ON charge measured at the reversal potential. Figure 7A shows superimposed ON gating and tail currents of α_1C+β₃, recorded without prepulses and after 50-ms repolarization to −90mV following a 200-ms prepulse to 60mV. Both gating and tail currents increased in magnitude after the prepulse. Some degree of facilitation of gating and ionic currents remained when the repolarization time was increased to 1s (Figure 7B). The insets in Figure 7AB show enlarged gating currents. The solid lines represent time integrals of the gating currents recorded in the absence of prepulse, while dashed lines are the time integrals of the gating currents recorded after the 200-ms prepulse to 60mV. Clearly, the prepulse was able to enhance the total charge moved at the beginning of the depolarization.

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Figure 7

Increase in charge movement and tail current after prepulse in α_1C+β₃, and effect of the α₂δ subunit. In oocytes expressing α_1C+β₃ and α_1C+β₃+α₂δ, gating and ionic tail currents were simultaneously recorded pulsing to the reversal potential. The external solution contained 10mM Ba²⁺. (A) and (B) show superimposed current traces recorded from the same oocyte expressing α_1C+β₃ before and after a 200-ms prepulse to 60mV. In (A) the time of repolarization between the prepulse and the test pulse was 50ms, while in (B) it was 1s. The ON gating currents are magnified and their time integrals are shown. Solid lines represent time integrals of the gating currents recorded in the absence of prepulse; dashed lines are the time integrals of the gating currents recorded after the prepulse. The arrows indicate the amplitude of the tail currents recorded in the absence (−prepulse) or after (+prepulse) the prepulse. Notice that an increase in charge movement after the prepulse is accompanied by an increase in the tail current. (C) and (D) are current traces recorded from the same oocyte expressing α_1C+β₃+α₂δ before and after a 200-ms prepulse to 60mV. The repolarization time between the prepulse and the test pulse was 50ms (C) and 1s (D). A small decrease in charge movement and a corresponding decrease in tail current occurred when the α₂δ subunit was coexpressed.

After the coexpression of the α₂δ subunit (α_1C+β₃+α₂δ), an equivalent pulse protocol failed to produce an increase of both gating and ionic currents (Figure 7CD). Instead, both the gating and ionic current (tail) amplitudes were slightly reduced when the repolarizing interpulse was 50ms long (Figure 7C). The reduction in charge movement was probably due to the occupancy of closed states nearer to the open state on the activation pathway, because the short repolarizing interpulse was not long enough to allow for the re-population of the deepest closed states. For longer repolarization times, the gating and ionic currents were identical with and without prepulses (Figure 7D).

The time course of the decaying phase of the gating current was not significantly modified by the prepulse. The gating current decays were fitted with a double-exponential function. In α_1C+β₃, τ_fast and τ_slow were, respectively, 0.42±0.04ms and 3.98±0.99ms without prepulse, and 0.41±0.06ms and 3.44±0.88ms after the prepulse. In α_1C+β₃+α₂δ, τ_fast and τ_slow were, respectively, 0.44±0.06ms and 5.39±1.84ms without prepulse, and 0.39±0.05ms, 5.39±1.84 after the prepulse. The fast component of the total charge was predominant (87% for α_1C+β₃ and 92% for α_1C+β₃+α₂δ).

In order to better characterize the effect of prepulses on gating currents, we blocked the ionic conductance by replacing Ba²⁺ in the external solution with 2mM Co²⁺ and 0.2mM La³⁺. Similar to the experiments for the facilitation of the ionic current, we used 200-ms prepulses to 80mV. In α_1C+β₃-expressing oocytes, the prepulse produced an increase of charge movement at all of the tested potentials. Fig. 8 shows gating current traces (solid lines) and their time integral (dashed lines) from an oocyte expressing α_1C+β₃, recorded by stepping to the indicated potential from −90mV holding potential in control condition (without prepulse, Figure 8A) and after a 200-ms prepulse to 80mV (Figure 8B). Both ON and OFF gating currents were larger after the prepulses (B). The increase in charge movement was transient and recovered after holding the oocytes at −90mV for ∼2min (Figure 8C). We constructed Q-V curves by integrating the ON gating current measured during depolarizations between −80 and +70. Above 70mV the isolation of the gating current could not be adequately maintained due to the contaminating outward ionic current. In oocytes expressing α_1C+β₃, the charge increased ∼20% after the prepulse (Figure 8D). The Q-V curves for control, prepulse, and recovery were simultaneously fitted by a single Boltzmann distribution (solid lines) with a half-activation potential (V_half)=−9.9mV and effective valence (z)=1.4. The normalized maximum charge displacement after the prepulse increased by ∼20% compared to the control. No significant changes in the kinetic properties and voltage dependence of gating current were detected after the prepulses (Figure 8E).

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Figure 8

Prepulse effect on gating current in α_1C+β_3.α_1C+β₃ Gating current traces during 30-ms depolarization to the indicated potentials are shown. The ionic current was blocked by external 2mM Co²⁺ and 0.2mM La³⁺. (A) Gating currents recorded in the absence of a positive prepulse. In (B) the depolarizing steps were preceded by a 200-ms prepulse to +80mV followed by a 50-ms repolarization to −90mV. (C) Same conditions as in (A); the protocol for the recovery was applied 2min after the prepulse experiment. Dashed lines represent the time integrals of the gating currents. (D) Averaged Q-V curves (Q_ON) obtained from gating currents recorded in control conditions (■), after a 200-ms prepulse to +80mV (●), and during the recovery (▴) (n=5 oocytes). Error bars represent SEM. Solid lines are the simultaneous fits to a single Boltzmann distribution. Parameters used for the fitting are shown in the plot. (E) Normalized Q-V curves (same as in D): control (●), prepulse (■), and recovery (▴).

Similarly to the ionic current, the gating current of oocytes expressing α_1C+β₃+α₂δ did not increase with the prepulse (Figure 9AB). Instead, gating current amplitudes recorded after the 200-ms prepulse to 80mV (Figure 9B) were slightly reduced in respect to the gating current recorded in the absence of prepulse (Figure 9A). The Q-V curves obtained by integrating the ON gating current showed a small reduction of the maximum charge displacement when the positive prepulse was applied (Figure 9D). This small reduction was fully recovered in <1s at −90mV.

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Figure 9

Gating currents do not potentiate in the presence of the α₂δ subunit. Representative traces of gating currents from an oocyte expressing α_1C+β₃+α₂δ are shown. Gating currents were recorded in external 2mM Co²⁺ and 0.2mM La³⁺ during 30-ms depolarization to the indicated potentials. (A) Gating currents recorded in the absence of a positive prepulse. In (B) the depolarizing steps were preceded by a 200-ms prepulse to +80mV with a time of repolarization of 50ms between the prepulse and the test pulse. (C) Same conditions as in (A); the protocol for the recovery was run 2min after the prepulse experiment. The dashed lines are the time integral of the gating current. (D) Averaged Q-V curves (Q_ON) obtained from gating currents recorded in control conditions (■), after a 200-ms prepulse to +80mV (●), and after the recovery from potentiation (▴) (n=4). Error bars represent SEM. Solid lines are the simultaneous fits to a single Boltzmann distribution, and the parameters used for the fitting are shown (E). Normalized Q-V curves (same as in D): control (●), prepulse (■), and recovery (▴).

As for α_1C+β₃, the normalized Q-V curves from α_1C+β₃+α₂δ were simultaneously fitted by a single Boltzmann function having V_half=−2.29mV, z=1.81, with a reduction of ∼7% in Q_max in the presence of the prepulse (Figure 9D). No significant changes in the voltage dependence were observed after the prepulse (Figure 9E).

The voltage-dependent facilitation of the L-type Ca²⁺ current has been observed in a variety of tissues and cell lines. Although several studies concluded that cAMP-dependent phosphorylation is not involved in the voltage-dependent facilitation of L-type current, in some cases it has been shown the opposite (for a review see Dolphin, 1996). Some authors reported that the enhancement of the current level via the activation of PKA was not observed in the Xenopus oocyte expression system, probably because of the high basal level of PKA activity in oocytes (Singer-Lahat et al).

The voltage-dependent facilitation described in the present work was not significantly affected by altering the kinase activity of the oocytes. Injection of the non-hydrolyzable cAMP analog Rp-cAMP (100nl, 0.2mM or 2mM, n=10), with and without BAPTA, 0.5 to 3h before voltage clamp recording, did not cause significant changes in the prepulse facilitation (data not shown). Similarly, the cAMP-dependent PK inhibitor (6–22 amide) injected 10min before recording (100nl, 20μM) did not prevent long-lasting facilitation (n=3, data not shown). This result supports the hypothesis that the long-lasting facilitation of α_1C+β₃ is due to a structural change induced by strong depolarizations that set the channel in a conformational state more responsive to voltage. In agreement with the recent work of Dai et al, prepulse facilitation does not seem to require cAMP-dependent phosphorylation.

Some aspects of prepulse facilitation in L-type calcium channels resemble the characteristic voltage-dependent block by the G-protein of N- and E-type Ca²⁺ channels. The binding of the G-protein βγ subunit to the Ca²⁺ channel is voltage-dependent. Strong depolarizations relieve the block, leading to an increase in open probability of the channels (Bean, 1989,Ikeda, 1996) resulting in facilitation. Although cardiac α_1C channels expressed in oocytes are not modulated by G-protein (Qin et al), it is possible that a mechanism similar to the one mentioned is responsible for the long-lasting voltage-dependent facilitation of α_1C+β₃. The tonic inhibition of an unidentified molecule would be relieved by the strong depolarizations, yielding to the facilitation: the extremely slow ON rate of the rebinding of the blocking particle could be the reason for the long-lasting characteristic of the facilitation.

The coexpression of the α₂δ subunit seems to prevent the voltage-dependent facilitation of α_1C+β₃. The analysis of the voltage dependence of the activation suggests that the channels expressed with the α₂δ subunits behave as if they were constitutively potentiated. In fact, the G-V curve of α_1C+β₃+α₂δ is shifted to the left and its voltage dependence strictly follows the voltage dependence of the α_1C+β₃ channels when they are potentiated by the prepulse. The positive prepulse has only a very small effect on the G-V curves of channels expressed with α₂δ. An appealing interpretation of this result is the lack of room for channel potentiation, because the activation curve is already fully shifted to the left on the voltage axis. However, it should also be taken into consideration that in α_1C+β₃, the limiting G_max with and without prepulse tends to maintain the same value. This could be due either to the fact that the limiting open probability cannot be further increased by the prepulse, or because of a rapidly developing facilitation at very positive potentials (i.e., +180mV) during the 25-ms test pulse itself. Then, the more positive part of the G-V curve (without prepulse) may be steeper because of the facilitation that develops during the positive pulses, thereby producing the shift.

The G-V curves were obtained from tail currents, which were completely abolished by replacing the external 10mM Ba²⁺ with 2mM Co²⁺ and 0.2mM La³⁺. However, although the outward current was reduced by Co²⁺ and La³⁺, it was not completely blocked, possibly because of both the release of the block by the depolarizations and the outward ionic flux. The block of the current was restored instantaneously by membrane repolarization, as suggested by the lack of tail currents (data not shown, previously discussed in Olcese et al). Although the outward current is mainly carried by the expressed Ca²⁺ channels (it is strictly dependent on the level of Ca²⁺ channel expression and displays kinetics that depend on the subunit compositions of the expressed channel), at extreme positive potentials it could be contaminated by a small endogenous current.

Because α₂δ enhances the voltage-dependent inactivation, it is reasonable to consider that the absence of facilitation in the presence of the α₂δ subunit may be due to a counteracting effect on the facilitation. However, if this were the case, the two processes, facilitation and inactivation, should develop and recover with identical time courses but opposite amplitudes, exactly canceling each other out, to generate a result as the one shown in Figure 2F. Although this situation is possible, we consider it very improbable. Also, as indicated by the tail envelope test for pulses to 80mV of increasing duration (from 25 to 200ms) performed in the same batch of oocytes, facilitation of α_1C+β₃ and inactivation of α_1C+β₃+α₂δ develop with different time constants and relative amplitudes (data not shown). Furthermore, even though the degree of inactivation varies among batches of oocytes, we never detected facilitation in α2δ-expressing oocytes even when inactivation was practically absent during the 200-ms prepulse.

In α_1C+β₃, the amount of movable charge increased after prepulses, suggesting a recruitment of a fraction of channels that are normally unable to gate during moderate depolarizations. Instead, as it was the case for the ionic current, also the charge potentiation was absent when α₂δ subunits were coexpressed. We were unable to estimate the amount of the ionic current facilitation deriving from a change in gating mode and the fraction from the recruitment of new channels. It is possible that the fraction of charge facilitated by the prepulse is totally uncoupled to the channel opening. In fact, if the facilitation were produced only by newly recruited channels, it would generate an activation curve (G-V) with the same voltage dependence and higher limiting conductance. Instead, the left shift of the G-V curve associated with the kinetics changes of facilitated channels suggests a modification of the activation pathway. Nevertheless, the new channels recruited by the prepulse may contribute to the overall potentiation.

The study of the effect of prepulse facilitation on gating current can be complicated by the change in voltage dependence of the charge movement occurring in inactivated channels. As in other voltage-dependent ion channels, slow inactivation produces a shift to the left on the voltage axis of the Q-V curve in L-type Ca²⁺ channels. However, the shift to more negative potentials, as described by Shirokov end collaborators, 1998, is produced by long depolarizations, and is further increased by coexpression of the α2δ subunit. The prepulses used in this study are short (200ms), and no changes in the voltage dependence of the Q-V curves were detected after the prepulse (Figs. 8E and Figure 9E).

It has been previously shown that the voltage-dependent facilitation of L-type Ca²⁺ channels is dependent on the type of β subunit coexpressed (Cens et al). Here we have shown that strong depolarizations facilitate both charge movement and ionic current in α_1C+β₃. Thus, voltage-dependent facilitation must increase both the number of channels that are able to produce gating current and the number of channels that can open in a normal voltage range. We have also shown the involvement of the α₂δ subunit in the facilitation, and its effect on ionic and gating current. A direct interaction between the α₂δ and β subunits is rather unlikely, as only five amino acids of the α₂δ subunit are intracellular, while the entire β subunit is known to be intracellular. The modulation exerted by the α₂δ subunit on α_1C could instead result from a direct interaction with the pore forming subunit (Gurnett et al).