Interaction via a Key Tryptophan in the I-II Linker of N-Type Calcium Channels Is Required for β1 But Not for Palmitoylated β2, Implicating an Additional Binding Site in the Regulation of Channel Voltage-Dependent Properties

Abstract

The Ca_Vβ subunits of voltage-gated calcium channels regulate these channels in several ways. Here we investigate the role of these auxiliary subunits in the expression of functional N-type channels at the plasma membrane and in the modulation by G-protein-coupled receptors of this neuronal channel. To do so, we mutated tryptophan 391 to an alanine within the α-interacting domain (AID) in the I-II linker of Ca_V2.2. We showed that the mutation W391 virtually abolishes the binding of Ca_Vβ1b and Ca_Vβ2a to the Ca_V2.2 I-II linker and strongly reduced current density and cell surface expression of both Ca_V2.2/α2δ-2/β1b and/β2a channels. When associated with Ca_Vβ1b, the W391A mutation also prevented the Ca_Vβ1b-mediated hyperpolarization of Ca_V2.2 channel activation and steady-state inactivation. However, the mutated Ca_V2.2W391A/β1b channels were still inhibited to a similar extent by activation of the D₂ dopamine receptor with the agonist quinpirole. Nevertheless, key hallmarks of G-protein modulation of N-type currents, such as slowed activation kinetics and prepulse facilitation, were not observed for the mutated channel. In contrast, Ca_Vβ2a was still able to completely modulate the biophysical properties of Ca_V2.2W391A channel and allow voltage-dependent G-protein modulation of Ca_V2.2W391A. Additional data suggest that the concentration of Ca_Vβ2a in the proximity of the channel is enhanced independently of its binding to the AID by its palmitoylation. This is essentially sufficient for all of the functional effects of Ca_Vβ2a, which may occur via a second lower-affinity binding site, except trafficking the channel to the plasma membrane, which requires interaction with the AID region.

Introduction

Voltage-gated calcium (Ca_V) channels play a major role in the physiology of excitable cells, particularly of neurons. Three families of voltage-gated calcium channels have been identified, Ca_V1-Ca_V3 (for review, see Ertel et al., 2000). The Ca_V1 class, L-type channels and the Ca_V2 class, non-L-type channels are both high-voltage-activated (HVA). These are heteromultimers composed of the pore-forming α1 subunit, associated with auxiliary Ca_Vβ and α2δ subunits (for review, see Catterall, 2000). The Ca_V2 calcium channels are inhibited by Gβγ dimers (Herlitze et al., 1996; Ikeda, 1996), which is the main mechanism of presynaptic inhibition by G-protein-coupled receptors. Ca_Vβ subunits are crucial for normal HVA channel function (for review, see Dolphin, 2003a), because they enhance expression of functional channels at the plasma membrane, modulate their biophysical properties, and promote the voltage dependence of modulation of Ca_V2.2 calcium channels by Gβγ dimers, although the mechanism involved remains unclear (Bichet et al., 2000; Meir et al., 2000; Canti et al., 2001). Gβγ dimers and Ca_Vβ subunits have been shown to bind to overlapping sites in the I-II loop of Ca_V2 channels, and they induce opposite effects on biophysical properties of the channels (for review, see Dolphin, 2003b), shifting channels between a willing and a reluctant state that requires large depolarizations to be opened (Bean, 1989). This observation led to the hypothesis that the binding of Gβγ dimers might dissociate the Ca_Vβ subunit from the I-II loop of the α1 subunit (Sandoz et al., 2004) and that dissociation of Ca_Vβ was the mechanism responsible for the inhibition observed. However, there is also evidence that Gβγ dimers do not displace Ca_Vβ subunits but alter the orientation of the subunit with respect to the α1 subunit (Hummer et al., 2003).

Here, we investigated the role of Ca_Vβ subunits in the plasma membrane expression and G-protein modulation of Ca_V2.2 calcium channels by mutating the tryptophan (W391) in the α1-interacting domain (AID) in the I-II loop of Ca_V2.2. This amino acid has been shown both by the study that identified the AID motif and the recent structural studies to be key to the interaction between Ca_Vβ subunits and the AID (Pragnell et al., 1994; Chen et al., 2004; Opatowsky et al., 2004; Van Petegem et al., 2004). This mutation prevents the enhancement of functional expression of Ca_V2.2 by Ca_Vβ1b and also prevents modulation of Ca_V2.2 by this subunit. In addition, although the G-protein modulation of Ca_V2.2W391A was present, it was not voltage dependent. In contrast, only the expression of the channel at the plasma membrane was affected when Ca_Vβ2a was coexpressed with this mutant channel, whereas all of the biophysical properties of the expressed Ca_V2.2W391A channels were still normally modulated by Ca_Vβ2a, including the voltage dependence of G-protein modulation. Our results further show that this was dependent on palmitoylation of Ca_Vβ2a and suggest that the effect of Ca_Vβ subunits on the voltage-dependent properties of the Ca_V2.2 channels occurs not only via the high-affinity I-II linker interaction but also via low-affinity interactions, presumably with other sites on the channel.

Materials and Methods

Materials. The cDNAs used in this study were Ca_V2.2 (D14157), Ca_Vβ1b (X61394), Ca_Vβ3 (M88751), Ca_Vβ2a(M88751), Ca_Vβ2a-β1b chimera (Olcese et al., 1994), α2δ-2 (Barclay et al., 2001), and D₂ dopamine receptor (X17458). The green fluorescent protein (GFP-mut3b, U73901) was used to identify transfected cells. All cDNAs were subcloned in pMT₂.

Construction, expression, and purification of proteins. Ca_V2.2W391A was generated by site-directed mutagenesis with primers for Ca_V2.2W391A.F (5′CGGGTACCTGGAGGCGATCTTCAAGGCTGAG) and Ca_V2.2W391A.R (5′CTCAGCCTTGAAGATCGCCTCCAGGTACCCG).

Ca_V2.2 I-II loop in pGEX2T (Bell et al., 2001) [glutathione S-transferase (GST)-Ca_V2.2 I-II loop] was modified by PCR using Pfu polymerase (Stratagene, Amsterdam, The Netherlands) with the oligonucleotide primers 5′-ATGCTGGCCGAGGAGGACAGGAATGCAGAG-3′ and 5′-GACTCATTCCTCCGCCTTGAAGATCCACTCCAGGTAC-3′, which truncate the Ca_V2.2 I-II loop after the region encoding the AID. In this way, a construct was generated encoding the first 40 residues of the Ca_V2.2 I-II loop, incorporating the AID, fused to GST to form GST-Ca_V2.2(357-397)WT. Mutagenesis of GST-Ca_V2.2(357-397)WT was performed as described above using the primers 5′-ATCTTCAAGGCGGAGGAATGAGTCATGCTGGCCG-3′ and 5′-AGCCTCCAGGTACCCGTTGAGCTCTCGCTCGATC-3′ to substitute an alanine for W391. GST-Ca_V2.2(357-397)WT, GST-Ca_V2.2(357-397)W391A, and GST were expressed and purified as described for GST-Ca_V2.2 I-II loop by Bell et al. (2001). N-Terminally His-tagged Ca_vβ2a (H6N-Ca_vβ2a) and C-terminally His-tagged Ca_vβ1b (Ca_vβ1b-H6C) were expressed and purified as described by Bell et al. (2001).

Surface plasmon resonance. Assays were performed using a BIAcore 2000 (Biacore, Uppsala, Sweden) at 25°C using running buffer (20 mm Na phosphate, pH 7.5, 500 mm NaCl, and 0.005% Tween 20). Anti-GST monoclonal antibodies (Amersham Biosciences, Little Chalfont, UK) were covalently attached to the surface of CM5 dextran sensor chips according to the instructions of the manufacturer to allow the reversible immobilization of GST-fusion proteins. Approximately equivalent molar quantities ∼350 resonance units (RU) of GST-Ca_V2.2(357-397)WT, ∼350 RU of GST-Ca_V2.2(357-397)W391A, and ∼300 RU of GST were immobilized on successive flow cells of a sensor chip. The flow rate of running buffer was 25 μl/min. H6N-Ca_vβ2a and Ca_vβ1b-H6C were dialyzed against running buffer and diluted to a series of concentrations. These samples were applied for 6 min over all flow cells, and each injection was followed by a similar dissociation phase. The sensor chip surface was regenerated between injections by the application of 35 μl of 20 mm glycine/HCl, pH 2.2, at 10 μl/min and the immobilization of fresh ligand.

Sensorgrams were processed using the BIAevaluation 3.0 software (Biacore). Sensorgrams recorded from the flow cells containing GST-Ca_V2.2(357-397)WT and GST-Ca_V2.2(357-397)W391A were corrected for passive refractive index changes and for nonspecific interactions by subtraction of the corresponding sensorgram recorded from the flow cell containing GST only. Sensorgrams were fitted by nonlinear regression using Prism 4 (GraphPad Software San Diego, CA). For each sensorgram, the first 120 s of the association phase and the dissociation phase were fitted to a single exponential to determine the observed association rate k_on(obs) and the dissociation rate k_off. The specific association rate, k_on, was calculated as k_on = (k_on(obs) - k_off)/[Ca_Vβ], and the dissociation constant K_D was calculated as K_D = k_off/k_on.

Cell culture and heterologous expression. The tsA-201 cells were cultured in a medium consisting of DMEM, 10% fetal bovine serum, and 1% nonessential amino acids. The cDNAs (all at 1 μg/μl) for Ca_Vα1 subunits, Ca_Vβ, α₂δ-2, D₂ dopamine receptor, and GFP (when used as a reporter of transfected cells) were mixed in a ratio of 3:2:2:2:0.4. The cells were transfected using Fugene6 (DNA/Fugene6 ratio of 2 μg in 3 μl; Roche Diagnostics, Lewes, UK). The tsA-201 cells were replated at low density on 35 mm tissue culture dishes on the day of recording.

Biotinylation. T75 flasks of transiently transfected tsA-201 cells were washed three times with 10 ml of PBS. Cells were incubated with 2 ml of PBS, pH 8.0, containing 800 μm EZ-link Sulfo NHS-SS-Biotin (Pierce, Rockford, IL) for 15 min at room temperature. The biotinylation reaction was terminated by addition of 10 ml of 100 mm glycine in PBS, and cells were collected by centrifugation (1000 × g) and washed an additional three times with 10 ml of PBS.

Cells were lysed by addition of 750 μl of lysis buffer (20 mm Tris, pH 7.4, 150 mm NaCl, 1 mm EDTA, and 1.5% Triton X-100) containing one Complete protease inhibitor tablet (Roche Diagnostics) per 10 ml, followed by brief sonication. After centrifugation at 20,000 × g for 20 min at 4°C, the protein concentration of the supernatant was determined using the BCA assay (Pierce). Biotinylated proteins were precipitated by incubating 500 μg of supernatant with 50 μl of streptavidin agarose beads (Pierce) for 3 h at room temperature. Precipitated proteins were washed four times with 1 ml of lysis buffer. Proteins were eluted from the beads by incubation with 100 mm dithiothreitol in lysis buffer for 30 min, followed by an equal volume of 2× SDS-PAGE sample buffer. Samples of supernatant and precipitated proteins were analyzed by SDS-PAGE on 4-12% Tris-glycine gels (Invitrogen, Paisley, UK) followed by Western blotting with anti Ca_V2.2 II-III linker antibodies (Raghib et al., 2001).

Western blot analysis. Samples (2.5-250 μg of protein) from tsA-201 whole-cell lysates [prepared as described for COS-7 cells by Raghib et al. (2001)] or from biotinylation experiments (see above) were separated by SDS-PAGE on 4-12% Tris-glycine gels and then transferred to polyvinylidene fluoride membranes. Immunodetection was performed with antibodies to the Ca_v2.2 II-III linker (Raghib et al., 2001) or β3 subunit (Canti et al., 2001) as described previously (Raghib et al., 2001).

Immunocytochemistry. Two days after transfection with Ca_Vβ2a or Ca_Vβ2aC3,4S, tsA-201 cells were fixed with 4% paraformaldehyde in PBS for 5 min at room temperature. For permeabilization, cells were incubated twice for 7 min in a 0.02% solution of Triton X-100 in Tris-buffered saline. For detection of Ca_Vβ2a or Ca_Vβ2aC3,4S, the primary antibody used was a rabbit anti-Ca_Vβ2a (462-600) at 0.2 μg/ml (Chien et al., 1995). The secondary antibody was an anti-rabbit IgG FITC conjugated (1:500; Sigma, St. Louis, MO). For nuclear staining, 4′,6-diamidino-2-phenylindole (DAPI) (300 nm; Molecular Probes, Eugene, OR) was applied. Samples were mounted in VectaShield (Vector Laboratories, Burlingame, CA) to reduce photobleaching and examined on a confocal laser scanning microscope (model LSM; Ziess, Oberkochen, Germany) using a 40× objective (1.3 numerical aperture) with constant photomultiplier settings.

Whole-cell electrophysiology. Whole-cell patch-clamp recordings were performed at room temperature (22-24°C). Only fluorescent cells expressing GFP were used for recording. The single cells were voltage clamped using an Axopatch 200B patch-clamp amplifier (Axon Instruments, Foster City, CA). The electrode potential was adjusted to give zero current between pipette and external solution before the cells were attached. The cell capacitance varied from 10 to 40 pF. Patch pipettes were filled with a solution containing the following (in mm): 140 Cs-aspartate, 5 EGTA, 2 MgCl₂, 0.1 CaCl₂, 2 K₂ATP, and 10 HEPES, titrated to pH 7.2 with CsOH (310 mOsm; with a resistance of 2-3 MΩ). The external solution contained the following (in mm): 150 tetraethylammonium bromide, 3 KCl, 1.0 NaHCO₃, 1.0 MgCl₂, 10 HEPES, 4 glucose, and 10 BaCl₂, pH adjusted to 7.4 with Tris base (320 mOsm). The pipette and cell capacitance as well as the series resistance were compensated by 80%. Leak and residual capacitance current were subtracted using a P/4 protocol. Data were filtered at 2 kHz and digitized at 5-10 kHz. The holding potential was -100 mV, and pulses were delivered every 10 s.

Data analysis and curve fitting. Current amplitude was measured 10 ms after the onset of the test pulse, and the average over a 2 ms period was calculated and used for subsequent analysis. The current density-voltage (I-V) relationships were fitted with a modified Boltzmann equation as follows: I = G_max × (V - V_rev)/(1 + exp(-(V - V_{50, act})/k)), where I is the current density (in picoamperes per picofarad), G_max is the maximum conductance (in nanosiemens per picofarad), V_rev is the reversal potential, V_{50, act} is the midpoint voltage for current activation, and k is the slope factor. Steady-state inactivation properties were measured by applying 5 s pulse from -120 to +20 mV in 10 mV increments, followed by a 11 ms repolarization to -100 mV before the 100 ms test pulse to +20 mV. Steady-state inactivation and activation data were fitted with a single Boltzmann equation of the following form: I/I_max = (A₁ - A₂)/[1 + exp((V - V_{50, inact})/k)] + A₂, where I_max is the maximal current, and V_{50, inact} is the half-maximal voltage for current inactivation. For the steady-state inactivation, A₁ and A₂ represent the proportion of inactivating and non-inactivating current, respectively. Inactivation kinetics of the currents were estimated by fitting the decaying part of the current traces with the following equation: I(t) = C + A × exp(-(t - t_o)/τ_inact), where t_o is zero time, C the fraction of non-inactivating current, A the relative amplitude of the exponential, and τ_inact is its time constant. Activation kinetics were estimated by fitting the activation phase of the current with either a single or a double exponential. Analysis was performed using pClamp6 and Origin 7. Data are expressed as mean ± SEM of the number of replicates (n). Error bars indicate SEs of multiple determinations if not otherwise mentioned. Statistical significance was analyzed using Student's paired and unpaired t test, and p < 0.05 was considered to be statistically significant.

Results

Mutation of W391A in the I-II linker of Ca_V2.2 disrupts both Ca_Vβ1b and Ca_Vβ2a subunit binding

The amino acid W391 in Ca_V2.2 (Fig 1A) is conserved in the AID sequence of all HVA calcium channels (for review, see Dolphin, 2003a; Richards et al., 2004) and has been described previously to be important for the binding of the Ca_Vβ ancillary subunits to HVA calcium channels (Pragnell et al., 1994; De Waard et al., 1996; Berrou et al., 2002). The recent structural analysis of the interaction of Ca_Vβ subunits with the Ca_V1.2 I-II linker showed that this residue is deeply embedded in the AID binding groove in Ca_Vβ (Chen et al., 2004; Opatowsky et al., 2004; Van Petegem et al., 2004). We first showed by surface plasmon resonance analysis that mutation of W391 to A in the AID of Ca_V2.2 prevents the binding of both Ca_Vβ1b and Ca_Vβ2a subunit to the I-II linker of Ca_V2.2 (Fig. 1B). In our experiments, GST-fusion proteins corresponding to the proximal I-II linker, including the AID of Ca_V2.2 or Ca_V2.2W391A, or GST alone as control, were immobilized via an anti-GST antibody to an individual flow cell of a CM5 dextran sensor chip. Ca_Vβ subunit solutions (25-200 nm) were perfused over all flow cells. No binding of the Ca_Vβ subunits to the control GST-fusion protein was detected (data not shown). Ca_Vβ1b and Ca_Vβ2a exhibited specific binding to the I-II linker of Ca_V2.2. From the data shown in Figure 1B, the dissociation constants (K_D) of Ca_Vβ1b and Ca_Vβ2a for the I-II loop of Ca_V2.2 were calculated to be 13.8 and 10.3 nm, respectively. Negligible binding of either Ca_Vβ subunit was detected to the mutated Ca_V2.2W391A I-II linker, and thus the K_D values could not be determined. We can only estimate that they must be at least 1000-fold less than for the I-II linker of the wild-type channel. From this assay, we show that the W391 present in the AID is crucial for the binding of Ca_Vβ subunits to the proximal Ca_V2.2 I-II linker, and that the affinity for β binding to this construct is very similar to that observed previously for the full-length I-II linker (Bell et al., 2001).

Figure 1.

The W391A mutation prevents the binding of Ca_Vβ1b and Ca_Vβ2a to Ca_V2.2 I-II linker. A, Representation of the Ca_V2.2 subunit that is composed of four domains of six transmembrane segments each. The I-II linker contains an 18 amino acid domain (AID) that interacts with Ca_Vβ subunits (dotted box). The sequence of the AID within Ca_V2.2 and Ca_V2.2W391A are given below, with W391 underlined and also showing the putative overlapping binding motif for Gβγ dimer binding. B, Representative Biacore sensorgrams showing interactions between Ca_Vβ1b (top) or Ca_Vβ2a (bottom) with I-II loop GST-fusion proteins from Ca_V2.2 (left) and Ca_V2.2W391A (right). The auxiliary Ca_Vβ subunits (25-200 nm as indicated) were applied during the time indicated by the bars.

Ca_Vβ subunit binding is essential for functional expression of N-type calcium channels at the plasma membrane

Coexpression of Ca_V2.2 or Ca_V2.2W391A with accessory Ca_Vβ subunits allowed us to compare the biophysical properties of the wild-type and mutated channels. The α2δ-2 subunit was additionally present in all conditions in this study. The first striking effect of the W391A mutation was a large reduction of the current density for the mutated channel, consistent with a role of Ca_Vβ binding to the I-II linker in trafficking Ca_V1 and Ca_V2 channels to the plasma membrane (Bichet et al., 2000) (Fig. 2A). The G_max determined from the I-V relationships for Ca_V2.2W391A with Ca_Vβ1b was significantly decreased by 81 ± 3% compared with the Ca_V2.2/β1b combination (Fig. 2A; Table 1). The reduction in G_max was similar (94 ± 0.5%) when the wild-type channel was expressed in the absence of Ca_Vβ (Fig. 2A; Table 1). Similarly, the G_max was decreased by 72 ± 6% for Ca_Vβ2a with Ca_V2.2W391A compared with the Ca_V2.2/β2a combination (Fig. 2A; Table 1). In cells transfected with Ca_V2.2W391A in the absence of a Ca_Vβ subunit, none of the GFP-positive cells expressed any current (n = 21), whereas at least 90% of the GFP-positive Ca_V2.2/β1b-transfected cells displayed a current. This strongly suggests that Ca_V2.2 requires interaction via the I-II linker with a Ca_Vβ to traffic to the plasma membrane.

Figure 2.

Role of Ca_Vβ subunits in plasma membrane expression of Ca_V2.2. A, Left, I-V relationships for Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ1b (filled circles, left; n = 18) or Ca_Vβ2a (diamonds, right; n = 13) or without a Ca_Vβ subunit (filled squares, left; n = 13) compared with I-V relationships for Ca_V2.2W391A/α2δ-2 coexpressed with Ca_Vβ1b (filled stars, left; n = 11) or Ca_Vβ2a (filled triangles, right; n = 14). The mean data are fitted with a modified Boltzmann function (see Materials and Methods), the V_{50, act} and G_max values of which are given in Table 1. Typical Ba²⁺ current traces at +20 mV (identified by the symbols used) are shown above the I-V relationships. Right, Mean current density at +20 mV for each of these combinations ± SEM. B, Cell surface expression of either Ca_V2.2 or Ca_V2.2WA391A expressed with α2δ-2, either without Ca_Vβ or with Ca_Vβ1b (left) or with Ca_Vβ2a (right). Total expression of Ca_V2.2 is shown by Western blot in the top row and biotinylated Ca_V2.2 in the bottom row. Cells were transfected with empty vector (lane 1), Ca_V2.2/α2δ-2/β1b (lane 2), Ca_V2.2W391A/α2δ-2/β1b (lane 3), Ca_V2.2/α2δ-2 (lane 4), Ca_V2.2/α2δ-2/β2a (lane 5), or Ca_V2.2W391A/α2δ-2/β2a (lane 6). Note that only the cell surface expression is affected similarly by the W391A mutation or by the absence of Ca_Vβ. Right, Histogram showing quantification of the mean amount of Ca_V2.2WA391A expressed at the plasma membrane, when coexpressed with either β1b or β2a, or Ca_V2.2 without Ca_Vβ, given as a percentage of the amount of Ca_V2.2 expressed with the relevant Ca_Vβ present under the same conditions. Data are mean ± SEM of four independent experiments. MW, Molecular weight. C, Western blot illustrating the endogenous expression of Ca_Vβ3 in tsA-201 cells. Gel loaded with 2.5 μg of protein prepared from cells transfected with Ca_Vβ3 (lane 1) compared with 2.5 or 250 μg of protein from cells transfected with an empty pMT2 vector (lanes 2, 3). An anti-β3 monoclonal antibody was used for immunoblotting.

Table 1.

Biophysical properties of Ca_v2.2 and Ca_v2.2W391A

	Maximum conductance (Gmax′ nS/pF)	Activation V50,act (I-V curves) (mV)	Activation V50,act (tail currents) (mV)	Inactivation V50,inact (mV)
No Cavb
Cav2.2	0.28 ± 0.01	+14.8 ± 1.5	+24.8 ± 1.7	−35.1 ± 2.8
	n = 13		n = 8**	n = 6**
Cav2.2W391A	No expression
	n = 21
Cavβ1b
Cav2.2	4.5 ± 0.2	+5.2 ± 1.5	+13.6 ± 0.7	−48.6 ± 2.2
	n = 18		n = 19	n = 11
Cav2.2W391A	0.8 ± 0.02	+12.8 ± 2.1	+23.4 ± 1.4	−35.3 ± 3.5
	n = 11**		n = 12**	n = 8**
Cavβ3
Cav2.2	2.9 ± 0.07	+7.7 ± 0.3	N/D	−60.8 ± 0.7
	n = 7			n = 6
Cav2.2W391A	0.46 ± 0.03	+12.2 ± 0.9	N/D	−32.3 ± 1.6
	n = 7**			n = 8**
Cavβ2a
Cav2.2	4.0 ± 0.09	+9.3 ± 0.2	+20.4 ± 1.5	+6.8 ± 2.3
	n = 13		n = 10	n = 17
Cav2.2W391A	1.2 ± 0.04	+8.37 ± 0.3	+21 ± 1.5	+6.2 ± 2.6
	n = 14*		n = 12	n = 18
Cavβ2aC3,4
Cav2.2	4.6 ± 0.2	−1 ± 0.5	+9 ± 2.6	−44.6 ± 0.4
	n = 8		n = 10	n = 22
Cav2.2W391A	1.7 ± 0.07	+7.1 ± 0.7	+22.5 ± 2.5	−34.5 ± 3.2
	n = 7*		n = 11**	n = 10*
Cavβ2a/β1b
Cav2.2	7.1 ± 0.3	+4.9 ± 0.5	+19.4 ± 1.5	−11.6 ± 2.5
	n = 11		n = 14	n = 17
Cav2.2W391A	1.9 ± 0.08	+5.6 ± 0.5	+21.6 ± 1.6	−17.5 ± 0.6
	n = 12**		n = 14	n = 17

Statistical significance of the indicated parameter was determined by unpaired t test, comparing data obtained for the wild-type Ca_v2.2 channels with Ca_v2.2W391A when coexpressed without or with Ca_vβ subunits as indicated. ^*p < 0.05; ^**p < 0.01.

We used a biotinylation assay to assess biochemically whether there were fewer channels present at the surface of the tsA-201 cells transfected with Ca_V2.2W391A and a Ca_Vβ subunit or when Ca_V2.2 was expressed without a Ca_Vβ subunit, compared with wild-type Ca_V2.2/Ca_Vβ combination. Whereas the total expression of Ca_V2.2W391A was identical to the expression of Ca_V2.2 transfected with or without a Ca_Vβ, the amount of biotinylated channels at the plasma membrane was clearly lower (Fig. 2B). The W391A mutation decreased by 62 ± 5 and 37 ± 3% (n = 4) the number of channels that were present at the plasma membrane when coexpressed with Ca_Vβ1b or Ca_Vβ2a, respectively. A similar diminution of the surface expression (by 52 ± 6%) was observed for the wild-type Ca_V2.2 when expressed without Ca_Vβ (Fig. 2B). Even when the binding of Ca_Vβ subunits to the proximal I-II linker, including the W391A AID, was negligible in vitro, some channels were still able to traffic to the plasma membrane. It is therefore possible that Ca_Vβ subunits can bind to other binding sites present, for example, in the distal I-II linker or on the N and C terminus of the channel as suggested previously (Cornet et al., 2002; Maltez et al., 2005).

Given the foregoing results, the reason for the presence of a low level of wild-type Ca_V2.2 current in the absence of expressed Ca_Vβ may therefore be the presence of an endogenous β subunit as described previously in Xenopus oocytes (Canti et al., 2001). This was examined by Western blotting using specific Ca_Vβ antibodies. We clearly detected an endogenously expressed Ca_Vβ3 in tsA-201 cells (Fig. 2C). This endogenous Ca_Vβ may be responsible for trafficking wild-type Ca_V2.2, allowing small currents to be recorded in cells transfected without Ca_Vβ subunits, whereas the much lower affinity of the Ca_V2.2W391A channel would preclude any interactions with the low level of endogenous Ca_Vβ.

Biophysical properties of Ca_V2.2W391A expressed with Ca_Vβ1b are similar to those of Ca_V2.2 expressed without Ca_Vβ

Ca_Vβ1b is known to hyperpolarize the activation and the steady-state inactivation of HVA calcium channels (for review, see Dolphin, 2003a). If the W391A mutation were effectively disrupting the binding of Ca_Vβ1b to the channel, the biophysical properties of Ca_V2.2W391A should be comparable with those of Ca_V2.2 expressed without any Ca_Vβ subunit. This was indeed the case, because tail current analysis showed the V_{50, act} to be depolarized by +9.8 and +11.2 mV, respectively, for Ca_V2.2W391A/β1b and Ca_V2.2 in the absence of Ca_Vβ compared with Ca_V2.2/β1b (Fig. 3A,B; Table 1), confirming the V_{50, act} estimates obtained from the I-V relationships (Table 1). This indicated that the W391A mutation abolished the effect of Ca_Vβ1b on the voltage dependence of activation of Ca_V2.2, such that it behaved like the wild-type channel expressed without a Ca_Vβ subunit.

Figure 3.

Biophysical properties of Ca_V2.2 and Ca_V2.2W391A coexpressed with Ca_Vβ1b. A, Representative current traces to illustrate current activation. Ba²⁺ tail currents were recorded after repolarizing to -50 mV after a 20 ms test pulse to between -40 and +80 mV from a holding potential of -100 mV. Top, Ca_V2.2/β1b; middle, Ca_V2.2 without any Ca_Vβ; bottom, Ca_V2.2W391A/β1b, all coexpressed with α2δ-2. B, Voltage dependence of activation of Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ1b (filled circles) or without any Ca_Vβ subunit (filled squares) or Ca_V2.2W391A/α2δ-2 expressed with Ca_Vβ1b (filled stars). The normalized data, obtained from recordings such as those shown in A, are plotted against the test pulse (n = 6-19). The mean data are fitted with a Boltzmann function, the V_{50, act} values of which are given in Table 1. C, Representative current traces (labeled as in A) to illustrate steady-state inactivation protocols. Inward Ba²⁺ currents were recorded after conditioning pulses of 5 s duration, applied from a holding potential of -100 mV in 10 mV steps between -110 and +30 mV, followed by a 50 ms test pulse to +20 mV. D, Voltage dependence of steady-state inactivation of Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ1b (filled circles) or without any Ca_Vβ subunit (filled squares) or Ca_V2.2W391A/α2δ-2 expressed with Ca_Vβ1b (filled stars). The normalized data obtained from recordings such as those shown in C are plotted against the conditioning potentials (n = 6-19). The mean data are fitted with a Boltzmann function, the V_50,inact values of which are given in Table 1. E, Left, Superposition of representative current traces for the subunit combinations indicated, recorded during an 800 ms depolarizing step to +20 mV, from a holding potential of -100 mV, normalized to the peak current. Right, Mean time constants of inactivation (τ_inact) obtained by fitting the decaying phase of the Ba²⁺ currents at +20 mV with a single exponential, for Ca_V2.2/α2δ-2/β1b (black bar; n = 25), Ca_V2.2/α2δ-2 (white bar; n = 25), and Ca_V2.2W391A/α2δ-2/β1b (gray bar; n = 14).

Another important feature of Ca_Vβ1, β3, and β4 subunits is that they hyperpolarize the steady-inactivation curves of Ca_V2.2 as well as other HVA calcium channels (Bogdanov et al., 2000). The potential for half-inactivation (V_{50, inact}) was -48.6 mV for Ca_V2.2 expressed with Ca_Vβ1b (Fig. 3C,D; Table 1). Again, in agreement with the assumption that the I-II linker of the Ca_V2.2W391A channel does not bind Ca_Vβ1b, we found a significant depolarizing shift of the steady-state inactivation for Ca_V2.2W391A/β1b [by +13 mV (Table 1)], whose steady-state inactivation curve is superimposed on that of Ca_V2.2 expressed without Ca_Vβ1b (Fig. 3C,D; Table 1). Ca_Vβ subunits are known to modulate not only the voltage dependence of the inactivation of calcium channels but also the kinetics of current decay (for review, see Dolphin, 2003a). The time constant of inactivation (τ_inact) of the Ca_V2.2W391A/β1b current at +20 mV (209.2 ± 14.5 ms; n = 14) was smaller than that of the wild-type Ca_V2.2/β1b combination (510.3 ± 51.6 ms; n = 25; p < 0.01), whereas the wild-type Ca_V2.2 expressed without a Ca_Vβ subunit showed intermediate inactivation (Fig. 3E). The faster inactivation kinetics for the Ca_V2.2W391A/β1b currents might be explained by the introduction of the mutation itself, because it has been shown previously that mutations in the AID are able to alter the inactivation kinetics (Dafi et al., 2004; Berrou et al., 2005).

Altogether, these results indicate that Ca_V2.2W391A is not regulated by Ca_Vβ1b in the plasma membrane. Furthermore, if, as our evidence suggests, Ca_V2.2 expressed without Ca_Vβ subunit is trafficked to the plasma membrane by endogenous Ca_Vβ, this Ca_Vβ does not regulate the channels once they have reached the membrane, as suggested previously (Canti et al., 2001).

Differential modulation of N-type channels by Ca_Vβ2a subunits

Despite the fact that the W391A mutation effectively decreased the expression of Ca_V2.2W391A channels at the plasma membrane in the presence of Ca_Vβ2a as well as Ca_Vβ1b, no shift of activation was observed of the I-V curves when Ca_Vβ2a was coexpressed with Ca_V2.2W391A compared with the wild-type Ca_V2.2/β2a combination (Fig. 2A). This was confirmed by analysis of tail currents. The V_{50, act} was equivalent when either Ca_V2.2 or Ca_V2.2W391A was coexpressed with Ca_Vβ2a (Fig. 4A,B; Table 1).

Figure 4.

Biophysical properties of Ca_V2.2 and Ca_V2.2WA391A coexpressed with Ca_Vβ2a. A, Representative current traces to illustrate current activation using the same protocols as described in the legend to Figure 3. Top, Ca_V2.2/β2a; bottom, Ca_V2.2W391A/β2a, all coexpressed with α2δ-2. B, Voltage dependence of activation for Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ2a (filled diamonds; n = 10) or Ca_V2.2W391A/α2δ-2 expressed with Ca_Vβ2a (filled triangles; n = 12). The normalized data obtained from recordings such as those shown in A are plotted against the test pulse. The mean data are fitted with a Boltzmann function, the V_{50, act} values of which are given in Table 1. C, Representative current traces to illustrate steady-state inactivation using the same protocols as described in the legend to Figure 3. Top, Ca_V2.2/β2a; bottom, Ca_V2.2W391A/β2a. D, Voltage dependence of steady-state inactivation for Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ2a (filled diamonds; n = 17) or Ca_V2.2W391A/α2δ-2 expressed with Ca_Vβ2a (filled triangles; n = 18). The normalized data obtained from recordings such as those shown in C are plotted against the conditioning potentials. The mean data are fitted with a Boltzmann function, the V_{50, inact} values of which are given in Table 1. The dotted line represents the fit for Ca_V2.2 without Ca_Vβ from Figure 3D. E, Left, Superposition of representative current traces for the subunit combinations indicated, recorded during an 800 ms depolarizing step to +20 mV, from a holding potential of -100 mV and normalized to the peak current. Right, Normalized residual I_Ba at 600 ms, for Ca_V2.2/α2δ-2/β2a (black bar; n = 15) and Ca_V2.2W391A/α2δ-2/β2a (gray bar; n = 14).

As expected, Ca_Vβ2a depolarized the V_{50, inact} of Ca_V2.2 by +42 ± 2 mV compared with Ca_V2.2 expressed without Ca_Vβ (Fig. 4C,D; Table 1). However, contrary to all expectations, we found that Ca_Vβ2a was also able to depolarize the steady-state inactivation properties of Ca_V2.2W391A to the same extent as for wild-type Ca_V2.2 (Fig. 4D; Table 1). Furthermore, Ca_Vβ2a also slowed the inactivation kinetics of Ca_V2.2W391A currents (Fig. 4E), abolishing the acceleration of the inactivation observed when Ca_V2.2W391A was coexpressed with Ca_Vβ1b (Fig. 3E), as noted previously for other mutations in the AID region (Dafi et al., 2004). The decay of the currents was <40% during the 800 ms depolarizing pulses and therefore could not be fitted with an exponential function, so we estimated the inactivation rate from the ratio of the current at 600 ms to that at the peak. The ratio at +20 mV was equivalent when Ca_Vβ2a was coexpressed with either Ca_V2.2 (0.76 ± 0.06) or the mutated channel (0.77 ± 0.08) (Fig. 4E).

In summary, despite the fact that the W391 mutation was able to diminish the trafficking of the Ca_V2.2W391A channels to the plasma membrane with all of the Ca_Vβ subunits examined and in contrast to the results obtained for Ca_Vβ1b, the Ca_V2.2W391A/β2a channels expressed at the cell surface were still modulated by Ca_Vβ2a.

Interaction with a Ca_Vβ subunit is essential for the voltage dependence of the modulation of Ca_V2.2 calcium channels by G-protein activation

To investigate the importance of the W391 residue for the modulation by G-proteins of Ca_V2.2 calcium channels, we coexpressed a D₂ dopamine receptor with the Ca_V2.2, Ca_Vβ1b, α2δ-2 combination and activated the receptor with a maximal concentration (100 nm) of the agonist quinpirole. Figure 5A shows representative currents obtained before (P1) and immediately after (P2) a 100 ms depolarizing prepulse to +100 mV, before and during application of quinpirole. The currents measured at +10 mV were inhibited by quinpirole by 64.2 ± 4% for the wild-type channel (Fig. 5A, B). The P2/P1 ratio obtained from traces such as those represented in Figure 5A reflects the voltage-dependent loss of inhibition. A value of 2.3 ± 0.1 (n = 18) was obtained for P2/P1 at +10 mV for the wild-type channel expressed with Ca_Vβ1b (Fig. 5C).

Figure 5.

G-protein modulation of Ca_V2.2 and Ca_V2.2W391A expressed with Ca_Vβ1b. Top, The pulse protocol used is depicted. A 100 ms test pulse (P1) from -30 to +60 mV was applied from a holding potential of -100 mV. After 800 ms repolarization to -100 mV, a 100 ms prepulse to +100 mV was applied. The cell was repolarized for 20 ms to -100 mV, and a second pulse (P2) identical to the first one was applied. A, D, Typical current traces obtained with this protocol are represented for Ca_V2.2 (top) and Ca_V2.2W391A (bottom) coexpressed with Ca_Vβ1b. In A, the D₂ dopamine receptor is coexpressed, and the top current traces (depicted by the open symbols) are in the presence of the agonist quinpirole (100 nm). In D, Gβ1γ2 are coexpressed, and typical traces for Ca_V2.2/β1b and Ca_V2.2W391A/β1b are shown. B, I-V curves for the calcium channel combinations are shown, obtained before (filled symbols) and during (open symbols) application of 100 nm quinpirole. Top, I-V curves for Ca_V2.2/α2δ-2 coexpressed with Ca_Vβ1b (circles; n = 18) Bottom, I-V curves for Ca_V2.2W391A/α2δ-2 coexpressed with Ca_Vβ1b (stars; n = 11) are represented. I-V curves are fitted with modified Boltzmann functions, the V_{50, act} and G_max parameters of which are given in Table 1. E, Top, I-V curves for Ba²⁺ currents during P1 for Ca_V2.2/α2δ-2/β1b coexpressed without (filled circles; n = 6) or with (open triangles; n = 9) Gβ1γ2. Bottom, For Ca_V2.2W391A/α2δ-2/β1b coexpressed without (stars; n = 6) or with (open squares; n = 8) Gβ1γ2. I-V curves obtained from currents recorded during P2 when Gβ1γ2 was coexpressed with Ca_V2.2/α2δ-2/β1b (filled triangles; n = 9) or Ca_V2.2W391A/α2δ-2/β1b (filled squares; n = 8) are also represented. All data were obtained in parallel on the same experimental days. C, Voltage-dependent facilitation was calculated by dividing the peak current value obtained in P2 by that obtained in P1 at the potentials of 0, +10, and +20 mV, for Ca_V2.2/α2δ-2 with Ca_Vβ1b (black bars; n = 18), Ca_V2.2W391A/α2δ-2 with Ca_Vβ1b (white bars; n = 11) after application of quinpirole, or when Gβ1γ2 were coexpressed with Ca_V2.2/β1b (gray bars; n = 9), Ca_V2.2W391A/β1b (hatched bars; n = 8). F, Typical current traces at +20 mV for R52, 54ACa_V2.2W391A (top) coexpressed with Ca_Vβ1b before (filled circles) and after (open circles) activation of the D₂ dopamine receptor. Corresponding I-V curves are represented in the bottom.

For the Ca_V2.2W391A/β1b currents, inhibition by quinpirole was similar, being 58.8 ± 5.2% at +10 mV (n = 11) (Fig. 5A, B). This inhibition was prevented by preincubating the cells for 16 h with 100 ng/ml pertussis toxin (PTX), as was inhibition of the wild-type currents (data not shown). However, the P2/P1 ratio was markedly diminished to 1.3 ± 0.1 at +10 mV (n = 11; p < 0.01) (Fig. 5C), demonstrating a lack of voltage-dependent loss of the G-protein modulation for the mutated channel. In agreement with this, when the Ca_V2.2W391A/β1b combination was coexpressed with Gβ1γ2, it resulted in tonic inhibition compared with controls in the absence of Gβ1γ2, but these currents exhibited no slowed activation and no prepulse facilitation, in contrast to the wild-type Ca_V2.2/β1b currents (Fig. 5C-E). Furthermore, although the V_50,act of the I-V relationship was depolarized by +12.9 mV for the wild-type channel when Gβ1γ2 were cotransfected, it was only depolarized by +4.5 mV for the Ca_V2.2W391A channels, in agreement with a reduced voltage dependence of the modulation by Gβγ of Ca_V2.2W391A channels. Moreover, two arginines (R52 and R54) present in the N terminus of Ca_V2.2 are essential for the modulation by Gβγ of Ca_V2.2 calcium channels (Canti et al., 1999). After mutation of these two amino acids to alanines in the N terminus of Ca_V2.2W391A, quinpirole no longer inhibited the currents. This shows that these residues, and therefore Gβγ, are involved in the voltage-independent inhibition of Ca_V2.2W391A induced by activation of the D₂ dopamine receptor by quinpirole (Fig. 5F).

To investigate whether these data were contaminated by the increased inactivation rate of the Ca_V2.2W391A/β1b combination compared with wild-type Ca_V2.2/β1b currents, we also examined the quinpirole-mediated inhibition of the wild-type and mutated channels expressed with Ca_Vβ3, which produces more inactivation of wild-type Ca_V2.2 than does Ca_Vβ1b (Fig. 6A). Unsurprisingly, when Ca_V2.2W391A was coexpressed with Ca_Vβ3, the G_max was dramatically reduced by 83 ± 4% compared with wild-type Ca_V2.2/β3 (Fig. 6A; Table 1). There was also a +4.5 mV shift of the V_{50, act} for Ca_V2.2W391A compared with wild-type Ca_V2.2/β3, and the Ca_Vβ3 subunit did not hyperpolarize the steady-state inactivation of Ca_V2.2W391A (Table 1). This suggests that, like Ca_Vβ1b, Ca_Vβ3 is not able to modulate the biophysical properties of Ca_V2.2W391A. However, quinpirole still inhibited Ca_V2.2W391A/β3 currents by 57.8 ± 11.8% at +10 mV (p < 0.01) (Fig. 6A). Furthermore, the voltage-dependent facilitation at +10 mV was greatly diminished from 2.5 ± 0.2 when Ca_Vβ3 was coexpressed with wild-type Ca_V2.2 to 1.3 ± 0.2 (n = 7; p < 0.01) for the Ca_V2.2W391A/β3 combination (Fig. 6B).

Figure 6.

The kinetics of inactivation do not contaminate the properties of G-protein modulation of Ca_V2.2 and Ca_V2.2W391A. A, Typical current traces obtained for Ca_V2.2/α2δ-2 and Ca_V2.2W391A/α2δ-2 coexpressed with Ca_Vβ3 and the D₂ dopamine receptor are represented (top). I-V curves obtained for these combinations before (filled symbols) and during (100 nm) application of quinpirole (open symbols) are also shown (bottom). B, Voltage-dependent facilitation for Ca_V2.2/α2δ-2 (black bars; n = 7) or Ca_V2.2W391A/α2δ-2 coexpressed with Ca_Vβ3 (white bars; n = 7) obtained from the P2/P1 ratio when the D₂ dopamine receptor was activated by 100 nm quinpirole. C, Facilitation rate of G-protein-modulated channels. The duration of the prepulse was increased from 0 to 200 ms. The P2/P1 facilitation ratios are given for each prepulse, for Ca_V2.2 (open circles; n = 16) and Ca_V2.2W391A (open stars; n = 17) coexpressed with Ca_Vβ1b. Data are fitted with a single exponential, the time constant (τ_facil) of which is given in Results.

Although the Ca_V2.2W391A currents exhibited faster inactivation kinetics than the wild-type currents, the decrease of facilitation observed for the Ca_V2.2W391A currents was not attributable to inactivation occurring during either P1 or the prepulse, because the facilitation was similar for currents formed from Ca_V2.2 with either Ca_Vβ1b or Ca_Vβ3, despite the fact that the latter showed accelerated inactivation kinetics (at +20 mV; τ_inact = 174.1 ± 17.1 ms; n = 7). In addition, at all prepulse durations from 5 to 200 ms, the P2/P1 ratio for the Ca_V2.2W391A/β1b currents remained markedly reduced compared with Ca_V2.2/β1b (Fig. 6C). Therefore, the inactivation occurring during the prepulse is not responsible for the apparent decrease in facilitation of Ca_V2.2W391A. Furthermore, the kinetics of facilitation could be fit to a single exponential, whose time constant (τ_facil) for Ca_V2.2/β1b was 9.0 ± 1.4 ms (Fig. 6C), whereas the small residual facilitation was much slower for the Ca_V2.2W391A/β1b combination (τ_facil = 16.7 ± 3.2 ms). These results indicate that Ca_Vβ accelerates the kinetics of facilitation as described previously (Canti et al., 2000), but also that the presence of a Ca_Vβ subunit, bound with high affinity to the I-II linker of Ca_V2.2, is essential for the voltage-dependent loss of inhibition, i.e., the ability of a +100 mV prepulse to remove Gβγ-mediated inhibition in P2.

A related characteristic of the G-protein modulation of Ca_V2 channels is that the kinetics of current activation are slowed. Activation of the D₂ dopamine receptor significantly slowed the activation kinetics of Ca_V2.2 channels coexpressed with Ca_Vβ1b, increasing the time constant of activation (τ_act) from 2.0 ms to a combination of a similar fast τ_act,fast (2.2 ms) and a much slower τ_act,slow of 23.4 ms representing 36.5% of the current (Figs. 5A, 7A). In contrast, the activation kinetics for Ca_V2.2W391A/β1b measured during a pulse to +10mV were unaffected by quinpirole. τ_act remained fast both before (2.1 ms) and during (2.1 ms) activation of the receptor (Figs. 5A, 7A).

Figure 7.

G-protein modulation of the kinetics and voltage dependence of activation of Ca_V2.2 and Ca_V2.2W391A coexpressed with β1b. A, The τ_act for currents recorded in P1 as described in Figure 5 before and during perfusion of quinpirole in cells transfected with Ca_V2.2/α2δ-2/Ca_Vβ1b (white and black bars, respectively; n = 18) and with Ca_V2.2W391A/α2δ-2/β1b (gray and hatched bars, respectively; n = 11). For Ca_V2.2/α2δ-2/Ca_Vβ1b in the presence of quinpirole, the data were fit by a double exponential with a fast τ_act and a slow τ_act, the percentage of each component being given above the bars. B, Activation curves derived from tail current amplitude measurements for Ca_V2.2/α2δ-2/β1b (left) and Ca_V2.2W391A/α2δ-2/β1b (right). Top, Typical tail current traces recorded after a test pulse to +80 mV before (filled symbols) and during (open symbols) application of 100 nm quinpirole. Bottom, Peak tail current density, before (filled symbols) and during (open symbols) application of quinpirole, for Ca_V2.2/α2δ-2/β1b (circles, left) or Ca_V2.2W391A/α2δ-2/β1b (stars, right). Data are the mean ± SEM of 12-19 cells, and the solid lines are Boltzmann functions fits, the parameters of which are given in Results.

Another hallmark of G-protein modulation is the depolarization of the voltage dependence of current activation. For the wild-type Ca_V2.2 coexpressed with Ca_Vβ1b, application of quinpirole induced a +17.5 mV shift of the V_{50, act}, from +14.6 ± 1.8 to +32.1 ± 2.5 mV (n = 19; p < 0.01) (Fig. 7B). This shift was reversible on washout of quinpirole (data not shown). There was no significant inhibition of tail current amplitude at +80 mV (13.5 ± 18.0%), indicating that all of the inhibition was entirely voltage dependent. In contrast, for Ca_V2.2W391A coexpressed with Ca_Vβ1b, the tail currents were still significantly inhibited by 55.8 ± 11.9% at +80 mV (p < 0.01) (Fig. 7B). However, the V_{50, act} for the residual Ca_V2.2W391A/β1b was still depolarized by application of quinpirole by +14.9 mV, from +27.5 ± 3.7 to +42.4 ± 1.9 mV (p < 0.01) (Fig. 7B), as was also true for Ca_V2.2 expressed without a Ca_Vβ subunit (data not shown), possibly indicating that Gβγ can still affect the voltage dependence of gating of Ca_V2.2W391A by shifting the channel to a reluctant state. Altogether, these results strongly suggest that the loss of ability of Ca_V2.2W391A to bind Ca_Vβ1b was accompanied by the almost complete loss of voltage dependence of the G-protein modulation of N-type calcium channels.

Ca_V2.2W391A coexpressed with Ca_Vβ2a shows voltage-dependent G-protein modulation

When Ca_Vβ2a was coexpressed with either Ca_V2.2 or Ca_V2.2W391A, quinpirole significantly inhibited the currents, by 64.4 ± 6.2% (p < 0.01) and 77.5 ± 6.1% (p < 0.01), respectively (Fig. 8A). Furthermore, in agreement with our evidence that Ca_V2.2W391A was still associated with Ca_Vβ2a (Fig. 4), we observed voltage-dependent facilitation of the Ca_V2.2W391A/β2a combination, equivalent to that observed for the wild-type Ca_V2.2/β2a (Fig. 8A,B). The P2/P1 ratio at +10 mV was 4.3 ± 0.6 for the wild-type channel and 3.7 ± 0.4 for Ca_V2.2W391A (Fig. 8B).

Figure 8.

G-protein modulation of the kinetics and voltage dependence of activation of Ca_V2.2 and Ca_V2.2W391A coexpressed with Ca_Vβ2a. A, The effect of quinpirole (100 nm) is compared on the Ca_V2.2/α2δ-2/β2a (left) and Ca_V2.2W391A/α2δ-2/β2a (right) combinations. Representative currents traces using the P1/P2 pulse protocol shown in Figure 5, obtained before (Con) and during (Quin) application of 100 nm quinpirole. B, The P2/P1 facilitation ratios for Ca_V2.2/α2δ-2/β2a (black bars; n = 13) or Ca_V2.2W391A/α2δ-2/β2a (white bars; n = 14) at potentials between 0 and +20 mV are given. C, The τ_act values for currents activated by +20 mV steps before and during application of quinpirole. Values are given for Ca_V2.2/α2δ-2/β2a in control conditions (white bars) and in the presence of quinpirole (black bars; n = 13) or for the Ca_V2.2W391A (gray and hatched bars, respectively; n = 14). In the presence of quinpirole, the data were fit by two exponentials, with the percentage of the total represented by each component given above the bar. D, Activation derived from normalized tail current density before (filled symbols) and during (open symbols) application of quinpirole (100 nm) for the subunit combination Ca_V2.2/α2δ-2/β2a (left, diamonds; n = 10) or Ca_V2.2W391A/α2δ-2/β2a (right, triangles; n = 12). The solid lines are Boltzmann function fits to the mean data, the parameters of which are given in Results.

For the Ca_V2.2/β2a combination, τ_act was slowed by application of quinpirole from 2.2 ± 0.1 ms to a combination of a similar τ_{act, fast} (2.2 ms) and a very slow τ_{act, slow} (>400 ms), representing 51% of the current at 100 ms (Fig. 8C). The activation kinetics of Ca_V2.2W391A were similarly slowed from a τ_act of 2.4 ± 0.1 ms to a combination of a similar τ_{act, fast} (2.4 ms) and a very slow τ_{act, slow} (>400 ms), representing 46% of the current at 100 ms (Fig. 8C). From tail currents recorded before and during activation of D₂ dopamine receptors, we observed an equivalent shift of V_{50, act} of +12.0 ± 1.7 and +15.6 ± 2.3 mV for Ca_V2.2 and the mutated channel, respectively (Fig. 8D). At +80 mV, the control tail current amplitude was not significantly greater that that in the presence of quinpirole for Ca_V2.2W391A/β2a (Fig. 8D), in contrast to the results obtained when Ca_Vβ1b was coexpressed with Ca_V2.2W391A. We can conclude that Ca_Vβ2a was still able to support voltage-dependent removal of the inhibition of Ca_V2.2W391A induced by activation of the D₂ dopamine receptors.

Palmitoylation is responsible for the ability of Ca_Vβ2a to regulate the voltage-dependent properties of Ca_V2.2W391A

A peculiarity of Ca_Vβ2a is that it contains two cysteines in its N terminus that are palmitoylated. Palmitoylation of these residues strongly modulates the biophysical properties of calcium channels associated with Ca_Vβ2a, particularly their inactivation (Olcese et al., 1994; Chien et al., 1996; Qin et al., 1998; Bogdanov et al., 2000; Hurley et al., 2000; Restituito et al., 2000). To investigate the role of this palmitoylation, we used a Ca_Vβ2a with the cysteines C3 and C4 mutated to serines (Ca_Vβ2aC3,4S) (Bogdanov et al., 2000) that is unable to incorporate palmitate (Qin et al., 1998). Coexpressing Ca_V2.2 with Ca_Vβ2aC3,4S resulted in a significant hyperpolarization of the tail current V_{50, act} by -11.4 mV compared with Ca_V2.2/β2a (Fig. 9A; Table 1). The effect of Ca_Vβ2aC3,4S was therefore similar to Ca_Vβ1b. In contrast, when Ca_Vβ2aC3,4S was coexpressed with Ca_V2.2W391A, the voltage dependence of activation of the channel did not show any hyperpolarizing shift compared with Ca_V2.2W391A/β2a (Fig. 9A; Table 1), suggesting that, when not palmitoylated, the Ca_Vβ2aC3,4S subunit was not able to modulate the Ca_V2.2W391A channel.

Figure 9.

Effect of palmitoylation of Ca_Vβ subunits on the modulation of Ca_V2.2 and Ca_V2.2W391A channels. A, Voltage dependence of activation of Ca_V2.2 (filled symbols) or Ca_V2.2W391A (open symbols) with either Ca_Vβ2aC3,4S (triangles) or Ca_Vβ2a-β1b chimera (pentagons) and α2δ-2. The solid lines are Boltzmann function fits to the mean data, the parameters of which are given in Results. B, Voltage dependence of steady-state inactivation of Ca_V2.2 (filled symbols) or Ca_V2.2W391A (open symbols) with either Ca_Vβ2aC3,4S (triangles) or Ca_Vβ2a-β1b chimera (pentagons) and α2δ-2. The solid lines are Boltzmann function fits to the mean data, the parameters of which are given in Results. C, Left, Representative Ba²⁺ current traces recorded during an 800 ms depolarizing step to +20 mV from a holding potential of -100 mV. Right, The normalized amount of residual current at 600 ms, obtained for Ca_V2.2 or Ca_V2.2W391A coexpressed with α2δ-2 and Ca_Vβ2aC3,4S (black and white bars, respectively; n = 18 for both) or the Ca_Vβ2a-β1b chimera (gray and hatched bars; n = 23 and n = 19, respectively). D, Left, The effect of quinpirole (100 nm) is compared for the subunit combinations Ca_V2.2W391A/α2δ-2/β2aC3,4S (top traces) and Ca_V2.2W391A/α2δ-2/β2a-β1b chimera (bottom traces). Representative currents traces using the P1/P2 pulse protocol shown in Figure 5, obtained before (Con) and during (Quin) application of 100 nm quinpirole. Right, The P2/P1 ratio obtained from such traces for Ca_V2.2 and Ca_V2.2W391A coexpressed with α2δ-2 and Ca_Vβ2aC3,4S (black and white bars; n = 7 and n = 8, respectively) or the Ca_Vβ2a-β1b chimera (gray and hatched bars; n = 12 and n = 11 respectively).

As described previously (Qin et al., 1998; Bogdanov et al., 2000; Hurley et al., 2000; Restituito et al., 2000), coexpressing this mutated Ca_Vβ2aC3,4S subunit with Ca_V2.2 significantly hyperpolarized the steady-state inactivation compared with wild-type Ca_Vβ2a, by -51 mV (Fig. 9B; Table 1), and accelerated the kinetics of inactivation (Fig. 9C). In contrast, the steady-state inactivation V_{50, inact} of the Ca_V2.2W391A channel coexpressed with Ca_Vβ2aC3,4S was significantly depolarized by >10 mV compared with the inactivation of the Ca_V2.2 channel in the presence of Ca_Vβ2aC3,4S (Fig. 9B; Table 1), suggesting a lack of modulation of Ca_V2.2W391A by Ca_Vβ2aC3,4S. As expected, currents recorded from cells expressing Ca_V2.2W391A with Ca_Vβ2aC3,4S also exhibited fast kinetics of inactivation, similar to those of this channel when coexpressed with Ca_Vβ1b (Fig. 9C).

We next determined whether the G-protein modulation of the wild-type Ca_V2.2/β2aC3,4S combination would be affected by the lack of palmitoylation of the Ca_Vβ subunit. The currents in this case were still inhibited by 58.9 ± 2.6% (n = 8) (Fig. 9D), and the P2/P1 facilitation ratio was 3.5 ± 0.4 at +10 mV, comparable with that for the wild-type channel coexpressed with Ca_Vβ2a (Fig. 9D). However, when this mutated Ca_Vβ2aC3,4S was coexpressed with the mutated Ca_V2.2W391A, although the currents were still significantly inhibited by activation of the D₂ dopamine receptor by 61.4 ± 6.9% (n = 7) (Fig. 9D), the P2/P1 ratio was significantly diminished to 1.2 ± 0.7 at +10 mV (Fig. 9D). These results demonstrate that, when the W391A mutation in the I-II linker of Ca_V2.2 is accompanied by the mutation of the two palmitoylation sites in the N terminus of Ca_Vβ2a, it abolishes the ability of Ca_Vβ2a to support the voltage dependence of G-protein modulation of the channel. The lack of palmitoylation is confirmed by the altered distribution when expressed in tsA-201 cells, from wild-type Ca_Vβ2a being predominantly membrane associated to diffuse expression throughout the cytoplasm for Ca_Vβ2aC3,4S (Fig. 10A).

Figure 10.

Proposed mechanism for G-protein modulation of Ca_V2.2 calcium channels and the effect of palmitoylation on the interaction of Ca_Vβ subunits with the I-II linker. A, Confocal immunofluorescent images of β2a and β2aC3,4S subunits expressed in tsA-201 cells, using an anti-β2 antibody (green). Nuclei are visualized with DAPI (blue). Scale bars, 20 μm. Note the membrane localization of the β2a subunit (left), whereas the β2aC3,4S subunit is localized to the cytoplasm (right). B, Model for the mechanism of action of Gβγ binding to the channel to inhibit its activity. The N terminus of Ca_V2.2 containing two arginines (R52, R54) constitutes a motif implicated in the inhibition by G-proteins, whereas the voltage-dependent facilitation (loss of inhibition induced by strong depolarization) requires a bound Ca_Vβ subunit on the I-II linker. C, The GK domain of Ca_Vβ subunits interacts with the AID, allowing a low-affinity binding of its SH3 domain elsewhere on the channel (e.g., to the I-II loop of Ca_V2.2) (Maltez et al., 2005) to modulate the properties of the channel. Palmitoylation of Ca_Vβ by anchoring the subunit to the plasma membrane allows this low-affinity interaction to occur when the W391 in the AID is mutated to disrupt its interaction with the GK domain.

To confirm that this property was inherent to the palmitoylation of the Ca_Vβ subunit, we used a chimera in which the N terminus of Ca_Vβ1b was swapped with that of Ca_Vβ2a to obtain a palmitoylatable Ca_Vβ1b (Olcese et al., 1994). The Ca_Vβ2a-β1b chimera was able to depolarize the V_50,act of Ca_V2.2W391A currents (Fig. 9A), similar to Ca_Vβ2a. The chimeric Ca_Vβ2a-β1b subunit also significantly depolarized the steady-state inactivation of both the wild-type Ca_V2.2 channel and Ca_V2.2W391A by +37 and +17.8 mV, respectively, compared with channels containing the Ca_Vβ1b subunit with Ca_V2.2 or Ca_V2.2W391A (Fig. 9B; Table 1). Furthermore, the chimeric Ca_Vβ2a-β1b subunit slowed the inactivation of the wild-type channel and, to a lesser extent, the mutated Ca_V2.2W391A (Fig. 9C). Moreover, when coexpressed with Ca_Vβ2a-β1b, Ca_V2.2W391A currents were modulated by G-protein activation in a voltage-dependent manner (Fig. 9D). Ca_V2.2W391A exhibited prepulse facilitation during application of quinpirole; the P2/P1 ratio in this case was 3.0 ± 0.2, which was comparable with the value obtained for the wild-type channel expressed with Ca_Vβ2a-β1b (3.4 ± 0.2) (Fig. 9D). Altogether, these results show that the W391 is crucial for the voltage-dependent effects of Ca_Vβ1b and Ca_Vβ3 but not for those of palmitoylated Ca_Vβ2a, because palmitoylation of Ca_Vβ can restore the voltage dependence of modulation of the mutated Ca_V2.2W391A channel.

Discussion

Ca_Vβ subunits are membrane-associated guanylate-kinase proteins characterized by guanylate kinase-like (GK) domain that binds to the AID motif in the I-II loop of HVA Ca_Va1 subunits (Fig. 1A) and an Src homology 3 domain (SH3) (Hanlon et al., 1999). The 18 amino acid AID motif contains a conserved tryptophan that is crucial for binding Ca_Vβ (Pragnell et al., 1994; Berrou et al., 2002). Recent structural data from three groups has provided detailed information about the interaction between the AID-Ca_Vβ complex and confirmed that this tryptophan is deeply embedded in the binding groove within the GK of Ca_Vβ (Chen et al., 2004; Opatowsky et al., 2004; Van Petegem et al., 2004).

Requirement of Ca_Vβ for plasma membrane expression of HVA calcium channel

One of the main effects of Ca_Vβ subunits on HVA calcium channels is to increase current density. However, the mechanism for this increase remains controversial, being attributed to increased trafficking (Bichet et al., 2000), increased maximum open probability (Neely et al., 2004), or both. Our data agree with previous studies showing biochemically that the amount of Ca_V1.2 channels in the plasma membrane was increased by Ca_Vβ subunits (Altier et al., 2002; Cohen et al., 2005). We show that fewer channels are present at the surface when no Ca_Vβ subunits were coexpressed or when the mutated Ca_V2.2W391A channels were cotransfected with a Ca_Vβ. It has been suggested that a Ca_Vβ bound to the I-II linker may mask an endoplasmic reticulum (ER) retention signal present in the I-II linker of HVA calcium channels and favor the trafficking of the channel to the cell surface (Bichet et al., 2000). If the trafficking of the channel were the only process affected by the lack of a Ca_Vβ bound to the I-II linker of Ca_V2.2, the reduction of the current density, as judged by the G_max, and the amount of Ca_V2.2 expressed at the plasma membrane would be the same. However, the G_max was more affected than the plasma membrane expression of Ca_V2.2 (81 and 62% reduction, respectively, when coexpressed with Ca_Vβ1b). The reason for this is likely to be that Ca_Vβ not only traffics the channel to the surface but also increases the maximum open probability of the channel, as suggested previously (Wakamori et al., 1999; Meir et al., 2000; Neely et al., 2004).

When no Ca_Vβ was coexpressed with wild-type Ca_V2.2 channels, small currents remained, whereas none of the GFP-positive cells transfected with the mutated Ca_V2.2W391A channel alone expressed any current. This suggests that the endogenous Ca_Vβ3 that we have identified in tsA-201 cells was responsible for trafficking some wild-type Ca_V2.2 to the plasma membrane, and that the markedly reduced affinity of the W391A mutated channel for Ca_Vβ subunits prevented interaction with the endogenous Ca_Vβ3 subunits and thus their trafficking to the plasma membrane. When a Ca_Vβ subunit was cotransfected with Ca_V2.2W391A channel, a small fraction of these channels were then trafficked to the plasma membrane, presumably because the overexpressed Ca_Vβ was able to either bind with low affinity to the mutated I-II linker of Ca_V2.2W391A or mask other ER retention signals that may be present on other parts of the channel (Cornet et al., 2002). Our results therefore provide very strong evidence that the binding of a Ca_Vβ auxiliary subunit to the channel is an essential requirement for functional expression of Ca_V2.2 at the plasma membrane.

Role of Ca_Vβ in voltage-dependent modulation of N-type calcium channels

Apart from their important role in trafficking, Ca_Vβ subunits also modify the biophysical properties of calcium channels (for review, see Dolphin, 2003a). Despite our conclusion that interaction with Ca_Vβ (either endogenous or expressed) is essential for trafficking to the plasma membrane, when Ca_V2.2 was expressed alone, its activation and inactivation properties were shifted to more positive potentials compared with those for Ca_V2.2 coexpressed with Ca_Vβ1b or Ca_Vβ3. The Ca_V2.2W391A/β1b or Ca_Vβ3 combinations showed similar biophysical properties to wild-type Ca_V2.2 channels expressed without a Ca_Vβ. Thus, although the channels required a Ca_Vβ to be trafficked to the plasma membrane, they were not modulated by the Ca_Vβ subunits when inserted in the plasma membrane, in agreement with our previous results suggesting two binding sites with differing affinities (Canti et al., 2001).

G-protein modulation of Ca_V2.2 calcium channels requires Ca_Vβ subunits to exhibit voltage dependence

Another unexpected feature of our results was that the Ca_V2.2W391A/β1b currents were still inhibited by D₂ dopamine receptor activation to the same extent as wild-type Ca_V2.2 channels, although there was no voltage-dependent relief of this inhibition. In our experiments, maximal inhibition of the Ca_V2.2W391A/β1b currents remained fast, occurring within <10 s, and was PTX sensitive. Furthermore, Ca_V2.2W391A/β1b currents were also tonically inhibited by Gβγ in a voltage-independent manner. We suggest that Gβγ dimers are involved both in the voltage-independent modulation of Ca_V2.2W391A/β1b currents and in the voltage-dependent modulation of the wild-type Ca_V2.2 currents.

Two models have been proposed for the regulation of HVA calcium channels by G-proteins. In the first, Gβγ dimers displace Ca_Vβ from the I-II loop, resulting in the reluctant state (Sandoz et al., 2004), whereas in the second model, both Gβγ and Ca_Vβ subunits would be able to bind to the channel at the same time (Meir et al., 2000; Hummer et al., 2003; Richards et al., 2004), with Gβγ binding producing the reluctant state. In the present study, the fact that activating the D₂ dopamine receptor still leads to a large inhibition of Ca_V2.2W391A currents also suggests that the Gβγ has direct effects on channel gating and does not simply displace Ca_Vβ subunits. Moreover, Van Petegem et al. (2004) speculated that, because the Gβγ-AID affinity is at least 10- to 20-fold weaker than the Ca_Vβ-AID affinity and a major conformational change would be required for Gβγ to displace Ca_Vβ, this was unlikely to occur (Van Petegem et al., 2004).

We showed previously that two arginines in the N terminus of Ca_V2 calcium channels are critical for G-protein modulation (Canti et al., 1999). By mutating these residues in the N terminus of Ca_V2.2W391A, we demonstrate that they are also critical for the voltage-independent inhibition of the Ca_V2.2W391A channel, whereas the voltage-dependent removal of this inhibition, or facilitation, involving the unbinding of Gβγ requires Ca_Vβ to be bound to the I-II linker of the channel (Fig. 10B), as suggested previously (Meir et al., 2000).

Palmitoylation of Ca_Vβ2a subunits allows its interaction with Ca_V2.2W391A

Unlike other Ca_Vβ subunits, palmitoylated Ca_Vβ2a depolarizes, rather than hyperpolarizes, the steady-state inactivation of HVA calcium channels (Jones et al., 1998; Restituito et al., 2000). This observation led to the conclusion that the I-II linker might constitute the inactivation gate of HVA calcium channels, with its mobility being reduced by the palmitoylation of Ca_Vβ2a anchoring it to the plasma membrane (Restituito et al., 2000). In our study, the Ca_V2.2W391A channels trafficked to the plasma membrane by Ca_Vβ2a remained modulated by this subunit, in contrast to Ca_Vβ1b, and our evidence shows clearly that this is attributable to the palmitoylation of Ca_Vβ2a. The mutated Ca_VβC3,4S subunit was not able to modulate the biophysical properties of the Ca_V2.2W391A channels, demonstrating that palmitoylation of Ca_Vβ2a helps to anchor the subunit to the channel. This hypothesis was confirmed by creating a palmitoylatable Ca_Vβ1b that was able to modulate the biophysical properties of the Ca_V2.2W391A channel.

As shown recently (Cohen et al., 2005; Maltez et al., 2005) and confirmed here, the AID-GK interaction is essential to traffic the channels to the plasma membrane. Our data suggest that either this interaction is not promoted by palmitoylation of Ca_Vβ2a or Ca_Vβ2a is not palmitoylated during the trafficking process, until the Ca_V2.2/β2a complex is inserted in the plasma membrane.

Short Ca_Vβ subunits that lack the GK domain retain the ability to modulate the open probability of Ca_V1.2, suggesting another low-affinity binding site on the Ca_V subunit for the SH3 domain (Cohen et al., 2005). An interaction of the SH3 with the I-II linker has been then demonstrated that modulates biophysical properties of calcium channels such as their inactivation (Maltez et al., 2005). The authors suggest that the role of the high-affinity GK-AID interaction is to increase the local concentration of Ca_Vβ and to promote lower-affinity interactions. In a similar way, palmitoylation of Ca_Vβ2a, promoting its membrane localization (Fig. 10A) and thus increasing its local concentration, may allow its binding to the Ca_V2.2W391A channel via interaction with low-affinity binding sites, such as the C and N termini or other sites on the I-II linker (Fig. 10C), that might be critical for the voltage-dependent properties of the channel, including the voltage dependence of G-protein modulation (Walker et al., 1999; Stephens et al., 2000; Maltez et al., 2005).

We also show here that, despite differentially affecting the biophysical properties of calcium channels, Ca_Vβ1b and Ca_Vβ2a subunits play the same role in the voltage dependence of G-protein modulation. They are necessary for voltage-dependent facilitation presumably via the second low-affinity site. Our results therefore show conclusively that Gβγ dimers have a direct inhibitory effect on Ca_V2.2 channel gating, and that Ca_Vβ subunits are required for the voltage-dependent removal of this inhibition.