# The Interaction between the I-II Loop and the III-IV Loop of Cav2.1 Contributes to Voltage-dependent Inactivation in a β-Dependent Manner

Sandrine Geib, Guillaume Sandoz, Véronique Cornet, Kamel Mabrouk, Odile

Fund-Saunier, Delphine Bichet, Michel Villaz, Toshinori Hoshi, Jean-Marc

Sabatier, Michel de Waard

► To cite this version:

Sandrine Geib, Guillaume Sandoz, Véronique Cornet, Kamel Mabrouk, Odile Fund-Saunier, et al.. The Interaction between the I-II Loop and the III-IV Loop of Cav2.1 Contributes to Voltage-dependent Inactivation in a β-Dependent Manner. Journal of Biological Chemistry, 2002, 277 (12), pp.10003-10013. 10.1074/jbc.M106231200. hal-03440327

HAL Id: hal-03440327

https://hal.science/hal-03440327

Submitted on 25 Nov 2021

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# The Interaction between the I-II Loop and the III-IV Loop of Ca~v~2.1 Contributes to Voltage-dependent Inactivation in a ~β~-Dependent Manner*

Received for publication, July 5, 2001, and in revised form, December 11, 2001
Published, JBC Papers in Press, January 14, 2002, DOI 10.1074/jbc.M106231200

Sandrine Geib§¶, Guillaume Sandoz§¶, Véronique Cornet‡, Kamel Mabrouk||, Odile Fund-Saunier§, Delphine Bichet§, Michel Villaz§, Toshinori Hoshi**‡‡, Jean-Marc Sabatier||, and Michel De Waard§ §§

From ‡INSERM Unité 464, Laboratoire de Neurobiologie des Canaux Ioniques, Faculté de Médecine Nord, Boulevard Pierre Dramard, 13916 Marseille Cedex 20, France, §Équipe Miate INSERM 99-31, Commissariat à l'Energie Atomique, Laboratoire Canaux Ioniques et Signalisation, Departement de Biologie Moléculaire et Structurale, 17 Rue des Martyrs, 38054 Grenoble Cedex 9, France, ||Laboratoire de Biochimie, CNRS UMR 6560, Faculté de Médecine Nord, Boulevard Pierre Dramard, 13916 Marseille Cedex 20, France, and the **Department of Physiology and Biophysics, University of Iowa, Iowa City, Iowa 52242

We have investigated the molecular mechanisms whereby the I-II loop controls voltage-dependent inactivation in P/Q calcium channels. We demonstrate that the I-II loop is localized in a central position to control calcium channel activity through the interaction with several cytoplasmic sequences; including the III-IV loop. Several experiments reveal the crucial role of the interaction between the I-II loop and the III-IV loop in channel inactivation. First, point mutations of two amino acid residues of the I-II loop of Ca~v~2.1 (Arg-387 or Glu-388) facilitate voltage-dependent inactivation. Second, overexpression of the III-IV loop, or injection of a peptide derived from this loop, produces a similar inactivation behavior than the mutated channels. Third, the III-IV peptide has no effect on channels mutated in the I-II loop. Thus, both point mutations and overexpression of the III-IV loop appear to act similarly on inactivation, by competing off the native interaction between the I-II and the III-IV loops of Ca~v~2.1. As they are known to affect inactivation, we also analyzed the effects of ~β~ subunits on these interactions. In experiments in which the ~β~_4 subunit is co-expressed, the III-IV peptide is no longer able to regulate channel inactivation. We conclude that (i) the contribution of the I-II loop to inactivation is partly mediated by an interaction with the III-IV loop and (ii) the ~β~ subunits partially control inactivation by modifying this interaction. These data provide novel insights into the mechanisms whereby the ~β~ subunit, the I-II loop, and the III-IV loop altogether can contribute to regulate inactivation in high voltage-activated calcium channels.

The influx of calcium through voltage-gated calcium channels controls a variety of cellular processes ranging from mem-

brane excitability and synaptic efficacy to gene expression. Both the amplitude and the duration of the calcium influx shape the spatio-temporal efficacy of calcium signaling. A tight control of both processes is needed to avoid long term increases in intracellular calcium levels, which are cytotoxic to neurons. Although the control of calcium entry can be achieved in several ways, inactivation of voltage-gated calcium channels appears to represent a key molecular process. For instance, inactivation is considered as a candidate mechanism for synaptic depression (1, 2). Inasmuch as there are several calcium channel types, there are also several inactivation behaviors. L-type calcium channels inactivate slowly, whereas the neuronal N-, P/Q-, and R-type channels inactivate faster. These fundamental differences are linked to the pore-forming Ca~v~ subunit, which contains the major molecular determinants for inactivation, although auxiliary subunits can play a regulatory function in this process.

Although significant advances have been made to the understanding of the mechanisms whereby inactivation proceeds in voltage-dependent Na⁺ and K⁺ channels, there are still major challenges lying ahead in the calcium channel field. The first challenge is a result of the ever increasing number of molecular determinants proposed to play a role in inactivation. A study by Zhang *et al.* (3) revealed that the S6 region of domain I is a critical determinant of voltage-dependent inactivation in Ca~v~2.1 and Ca~v~2.3 channels. Since that pioneering work, several other sequences have been implicated in inactivation, including the I-II linker domain (4), the proximal carboxyl terminus (5), and the S6 regions in domains II, III, and IV (6–8). In the midst of this apparent complexity, and contrary to all expectations, it was found that voltage and calcium may use the same molecular determinants to inactivate calcium channels (9). This basic observation points to the possibility that there may be an elusive mechanism whereby the various structural elements identified so far are coordinated to produce inactivation in calcium channels. A further challenge in solving the issue of calcium channel inactivation stems from a still greater level of complexity that is introduced by ~β~ subunit regulation. Clearly, these auxiliary subunits possess an enormous potential for *in vivo* tuning of channel behavior (2, 10). This behavior is expected because ~β~ subunit can bind to several Ca~v~ structures: the I-II loop (11), the amino terminus (12, 13), and the carboxyl terminus (14). The most interesting candidate sequence to investigate in terms of ~β~-dependent inactivation

<footnote>* This work was supported in part by grants from INSERM and Commissariat à l'Energie Atomique. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.</footnote>

<footnote>¶ Both authors contributed equally to this work.</footnote>

<footnote>‡‡ Present address: Department of Physiology, 3700 Hamilton Walk, University of Pennsylvania, Philadelphia, PA 19104.</footnote>

<footnote>§§ Supported by an Action Concertée Incitative grant from the French Research Ministry. To whom correspondence should be addressed. Tel.: 33-4-38786813; Fax: 33-4-38785041; E-mail: mdewaard@cea.fr.</footnote>
appears to be the I-II loop. First, it contains the α₁ interaction domain (AID),<fnref n="1" /> which is the only Caᵥ sequence identified that is able to bind all β subunit isoforms (11). For some β subunits (i.e. β₃), it represents even the only interacting sequence (12, 15, 16). Second, this sequence seems to have a central role in many other β regulatory functions (14, 17). Third, deletions in β sequence, upstream and downstream of the β interaction domain, that interacts with the I-II loop, produce either faster or slower inactivation (18), presumably by altering the regulatory input of β subunit at its main interaction site. In support of such an interpretation, Qin and collaborators (19) found that the intermediate splicing region of β subunits, located immediately upstream of the β interaction domain site, is a critical β subunit determinant for inactivation. Overall, these data indicate that the I-II loop plays a crucial role in inactivation and that it represents a critical β-dependent element of this process.

Here, we have investigated some of the β-dependent mechanisms whereby the I-II loop controls inactivation in voltage-dependent P/Q calcium channels. We demonstrate that two nonconserved residues of Caᵥ2.1 (Arg-387 and Glu-388; Ref. 20) are required for the control of channel inactivation. This effect is a result of their implication in the interaction between the I-II and III-IV loops of the channel. Accordingly, a peptide derived from the III-IV loop potently modulates the voltage-dependent inactivation of Caᵥ2.1. Co-expression of the β subunit prevents this effect suggesting that the β subunit, two sites of mutations in the I-II loop, and the III-IV peptide contribute to inactivation by coordinated interactions. We propose that the β subunit regulates voltage-dependent inactivation by neutralizing the contribution of the III-IV loop to this process.

## EXPERIMENTAL PROCEDURES

**Peptide Synthesis**—Stepwise elongation of the 40-mer (III-IV loop from Ser-1535 to Ser-1574 of Caᵥ2.1; amidated form) was carried out on 0.35 mmol of Fmoc amide resin (1% cross-linked; 0.62 meq of amino group/g) using a peptide synthesizer (model 433A, Applied Biosystems Inc.). Trifunctional amino acids were side-chain protected as follows: trityl for Cys, His, and Asn; *t*-butyl for Ser and Asp; *t*-butyloxycarbonyl for Lys; and pentamethylchroman for Arg. Each coupling cycle comprised: (i) unblocking of the α-amino group by piperidine (18 and 20% in N-methylpyrrolidinone (NMP) for 3 and 8 min); washings with NMP (5 × 1 min); (ii) coupling of the Fmoc amino acids (1 mmol) as their hydroxybenzotriazole active esters in NMP. The peptide resin was treated 2 h at 25 °C with trifluoroacetic acid containing 5% thioanisole, 5% ethanediithol in a final volume of 10 ml/g of peptide resin. The peptide was precipitated and washed twice by adding cold diethyl ether. The resulting crude peptide was pelleted by centrifugation (3000 × g; 10 min), and the crude pellets were dissolved in H₂O and freeze-dried. The peptide (50-mg batches) was purified by preparative reversed-phase HPLC (PerkinElmer Life Sciences, ODS 20 µm, 100 × 10 mm) using a 2-h linear gradient from 0 to 50% acetonitrile in 0.1% trifluoroacetic acid/H₂O (buffer A) and 70% acetonitrile in H₂O containing 0.1% trifluoroacetic acid (buffer B) at a flow rate of 6 ml/min, with UV detection at 230 nm. Fractions were collected and analyzed by analytical HPLC. Fractions containing the purified peptide (>99% homogeneity) were pooled and lyophilized. The chemical identity of the synthetic peptide was assessed by amino acid analysis after acid hydrolysis (6 N HCl, 1% (mass/volume) phenol, 20 h, 120 °C); and mass determination by matrix-assisted laser desorption ionization-time of flight mass spectrometry.

**Plasmid Constructions**—AID chimeras were produced by a two-step polymerase chain reaction (PCR) procedure. First, two independent PCR reactions were carried out in which the cDNA sequences (that encode amino- and carboxyl-terminal sequences around AID of the full-length I-II<sub>A</sub> loop) were amplified with the use of chimera AID oligonucleotide sequences.

Second, a final PCR reaction was carried out after annealing of the two initial PCR products by their AID sequences. The final PCR product was subcloned in pGEX2TK vector by the use of BamHI and EcoRI sites that were present in the most extreme 5' and 3'-oligonucleotides. The following internal chimeric oligonucleotides were used: 1) for AID<sub>S</sub> chimera, forward: 5'-CGTGATCCAGCTCATGAGCCCGAAGGTGCCTCCTCCAGCTGCTGCTGCCGCCGCAG-3', reverse: 5'-CTTCGGGGCTACATGAGCTGGATCACGCAGGGCGAGGAGGTGATCCTCGCAGAGGAC-3'; 2) for AID<sub>B</sub> chimera, forward: 5'-GAAGATCCACTCCAAGTACCCATTCAGTTCTCGCTCAATCTGCTGTCGCCGCCGCAG-3', reverse: 5'-CTGAATGGGTACTTGGAGTGGATTCACAAGGCGGAGGAGGTGATCCTCGCAGAGGAC-3'; and 3) for AID<sub>C/D</sub> chimera, forward: 5'-AGTGATCCAGTCCAGGTAGCCTTTGAGGTGCCTCTTCCAGCTGCTGCTGCCGCCGCAG-5', reverse: 5'-CTCAAGGCTACCTGGACTGGATCACTCAGGCAGAAGAGGTGATCCTCGCAGAGGAC-3'. The extreme 5' and 3' oligonucleotides were derived from the Caᵥ2.1 cDNA sequence (GenBank accession no. X57477): forward, base pairs 1374-1415; and reverse, base pairs 1709-1754. Mutations in the AID<sub>A</sub> cDNA sequence were performed using the QuikChange™ site-directed mutagenesis kit (Stratagene). Mutations were performed directly on the wild-type I-II<sub>A</sub> loop itself subcloned in the BamHI and EcoRI sites of pGEX2TK vector. Mutagenic primers extended from Caᵥ2.1 base pairs: 1428-1466 (I385L), 1434-1472 (R387E), 1437-1475 (E388D), 1443-1481 (N390K), 1452-1489 (M393L), 1455-1491 (E394D), 1464-1502 (S397T), and 1465-1507 (K398Q). Next, the 1416-1723 BsmI fragment encoding the wild-type AID motif in Caᵥ2.1 (pSP72 vector) was replaced by chimeric and mutated AID sequences. To facilitate cloning, the third BsmI site in the 3' noncoding region was deleted by digestion of the BamHI-EcoRI fragments of the noncoding region, followed by recircularization of the plasmid. The chimeric and mutated regions were sequenced in both directions on an Automated Sequencer (ABI PRISM model 310, PerkinElmer Life Sciences). The pcDNA3.1/His-C (Invitrogen) construct containing the sequence of the I-II<sub>A</sub> loop was as described previously (17). For experiments with CD8, we used the α chain of the human CD8 clone, in which the cDNA sequence corresponding to its cytoplasmic part was replaced by Caᵥ2.1 cDNA sequences. The following CD8 constructs were made: 1) CD8-stop with no cytoplasmic sequence, except a Myc tag (CD8 sequence subcloned in the EcoRI and BamHI sites of pcDNA3.1(-)/Myc-His A (Invitrogen) with forward primer: 5'-CCGGAATTCACCATGGCCTTACCAGTGACCGCCTTGCTCCTGCCGCTG-3' and reverse primer: 5'-CGCGATCCCCCTGTGGTTGCAGTAAAGGGTGATAACCAGTGACAGCAG-3'); 2) CD8-Nt<sub>A</sub> (amino acids 1-98), subcloned as previously described for the I-II<sub>A</sub> loop in the BamHI and EcoRI sites of pcDNA3-CD8-βARK-Myc vector (17) after removing the β-adrenergic receptor kinase sequence (Caᵥ2.1 base pairs 294-317 (forward) and 567-587 (reverse)); 3) CD8-II-III<sub>A</sub> (amino acids 714-1254) again between CD8 and Myc of pcDNA3-CD8-Myc (base pairs 2436-2466 (forward) and 4023-4052 (reverse)); 4) CD8-III-IV<sub>A</sub> (amino acids 1520-1575 with primers containing base pairs 4854-4879 (forward) and 4993-5018 (reverse); 5) CD8-Ct1<sub>A</sub> (amino acids 1821-2196) with primers containing base pairs: 5754-5792 (forward) and 6861-6881 (reverse); and 6) CD8-Ct2<sub>A</sub> (amino acids 2106-2423) with primers containing base pairs 6607-6630 (forward) and 7545-7562 (reverse).

**Electrophysiological Recordings**—Stage V or VI oocytes, from Xenopus laevis frogs, were microinjected with a nanoliter injector (WPI) using 40–50 nl of various cRNA mixtures (0.3 µg/µl wild-type, chimera or mutant Caᵥ2.1 ± 0.1 µg/µl β₄ subunit or various αCD8 chimeras; CD8-NTₐ, CD8-I-IIₐ, CD8-II-IIIₐ, CD8-III-IVₐ, CD8-CT1ₐ, or CD8-CT2ₐ). Some cells were also coinjected with a mixture of wild-type or mutant Caᵥ2.1 cRNA and III-IVₐ or control peptide. The final concentration of the peptides reached in the cells was 10 µM. Cells were incubated in defined nutrient oocyte medium (21) for 4–7 days prior to recordings. These cells could be kept for up to 3 weeks in these conditions. Two-electrode voltage clamp was performed with a GeneClamp amplifier (Axon Instruments, Foster City, CA). The extracellular solution used was as follows (in mM): Ba(OH)₂ 40, NaOH 50, KCl 3, HEPES 10, niflumic acid 0.5, pH 7.4, with methanesulfonic acid. Electrodes filled with 140 mM KCl, 10 mM EGTA, and HEPES (pH 7.2) had resistances comprising between 0.5 and 1 megohm. Recordings were filtered at 2 kHz and sampled at 5–10 kHz. Leak subtraction was performed on-line by a P/4 protocol. Voltage-dependent activation curves were obtained by measuring the peak current amplitude after cell depolarization from a holding potential of –90 mV. Pulse frequency was every 15 s. For steady-state inactivation curves, cells were depolarized to 20 mV from various holding potentials. New holding potentials were maintained at least 30 s before pulse application. Data were analyzed using pClamp version 6.02 software (Axon Instruments). The

<footnote n="1">¹ The abbreviations used are: AID, α₁ interaction domain; GST, glutathione S-transferase; Fmoc, N-(9-fluorenyl)methoxycarbonyl; NMP, N-methylpyrrolidinone; HPLC, high performance liquid chromatography; HIT, half-inactivation time; HIP, half-inactivation potential; HAP, half-activation potential; F, fahrad(s).</footnote>
chimeras expressed a variable number and proportion of kinetic components precluding a detailed kinetic analysis for a clear comparison in inactivation kinetics. For these reasons, we chose to use half-inactivation time (HIT) values instead of time constants.

*Metabolic Labeling and CD8 Immunoprecipitation*—Metabolic labeling was performed by coinjecting 0.2 µCi of [³⁵S]methionine per oocyte with the cRNA coding for the various CD8-Ca<sub>v</sub>2.1 proteins. After injection, the oocytes were maintained 24 h in Barth's medium (in mM: 88 NaCl, 1 KCl, 0.82 MgSO<sub>4</sub>, 0.33 Ca(NO<sub>3</sub>)<sub>2</sub>, 0.41 CaCl<sub>2</sub>, 2.4 NaHCO<sub>3</sub>, 15 HEPES, pH 7.4, NaOH). Next, 30 oocytes/condition were resuspended on ice in 1 ml of Buffer A (50 mM Tris, 1% Triton X-100, pH 6.8) supplemented with a protease inhibitor mixture (Complete™, Roche Molecular Biochemicals) and homogenized with a 1-ml Potter homogenizer (Bioblock). The homogenate was incubated at 4 °C for 30 min under agitation, and then centrifuged 20 min at 10,000 × g (AM2.19 rotor, Jouan MR22i). The supernatant was collected with a Pasteur pipette, care being taken to avoid the lipid layer. The ³⁵S-CD8-Ca<sub>v</sub>2.1-Myc proteins were then immunoprecipitated for 3 h at 4 °C with 4 µg of anti-Myc IgG coupled to protein A-Sepharose. Immunoprecipitated proteins were loaded on 9–15% SDS-PAGE gel, and the gel was dried and exposed for autoradiography.

*In Vitro Translation*—³⁵S-Labeled probes were synthesized by coupled *in vitro* transcription and translation using the TNT™ system (Promega). Nonincorporated [³⁵S]methionine was removed by purification on a PD10 column (Amersham Biosciences, Inc.). Translated proteins were analyzed by loading them onto 5–15% SDS-PAGE gels.

*Binding of GST Fusion Proteins*—Fusion proteins were purified as previously reported (22). Excess glutathione was removed, first, by the use of a PD10 column, second by dialyzing against PBS buffer, pH 7.4, at 4 °C (Spectrum, 8000-Da molecular mass cut-off). Purified GST fusion proteins were coupled to glutathione-agarose beads by a 30-min incubation in buffer A: Tris-buffered saline (25 mM Tris, 150 mM NaCl (pH 7.4)) plus 0.1% Triton X-100. Residual unbound fusion proteins were removed by a first wash with buffer A. Binding of ³⁵S-I-II<sub>A</sub> loop was initiated by the addition of 1–2 µl of the translation product in a 0.5-ml volume of beads/buffer A. Estimated fusion protein concentration was 1 µM. The mixture was incubated overnight at 4 °C. Beads were then washed four times with 1 ml of buffer A, and the bound radioactive protein analyzed by SDS-PAGE and autoradiography. Non-specific binding is determined as the radioactivity associated to GST-glutathione-agarose beads. For binding of ³⁵S-β₄ onto the chimeric GST-I-II<sub>A</sub> loops, crude BL21 bacterial extracts were used, incubated with glutathione-agarose beads, unbound material washed with Tris-buffered saline, and binding performed with *in vitro* translated β protein.

## RESULTS

*Structural Differences between AID Sequences and Potential Importance in β Regulation of Inactivation*—Several published observations demonstrate that non-L-type and L-type channels differ in the regulation of their inactivation by β subunits (14). Although β subunits appear to affect the rate of L-type channel inactivation, steady-state inactivation seems to be less dependent on β subunit coexpression than non-L-type channels (9, 18, 23–24). These data suggest that there are some intrinsic molecular determinants differing between L-type and non-L-type Ca<sub>v</sub> channels in the control of inactivation by β subunits. As the I-II loop contains the main β binding site, we hypothesized that part of the differences in β regulation of inactivation between these two groups of calcium channels may be attributed to β binding sites. This hypothesis is supported by experimental evidence illustrating that deletions, mutations, or splicing within the I-II loop all modify the kinetics of inactivation (4, 17). A close examination of the amino acid residues that constitute the AID sequence of each Ca<sub>v</sub> channel reveals some interesting differences among L- and non-L-type channels (Fig. 1A). Alignment of the AID sequences illustrates that residue conservation within each subfamily of calcium channels (L-versus non-L-type) is much higher than previously recognized. AID<sub>C</sub> is identical to AID<sub>D</sub> (both are thereafter referred to as AID<sub>C/D</sub>), whereas AID<sub>S</sub> differs by only two residues from AID<sub>C</sub> and AID<sub>D</sub>. Additionally, AID<sub>A</sub>, AID<sub>B</sub>, and AID<sub>E</sub> differ by only three residues at approximately the same location at which

<figure><img src="image_2.png" /></figure>

<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /></figure>

**FIG. 1.** Membrane expression levels and binding of the β₄ subunit to AID Ca₂.1 chimera channels. A, schematic diagram illustrating the substitution of AID<sub>A</sub> by homologous non-L- and L-type AID sequences. Sequence alignment shows the homologies between the various AID sequences. Residues in bold correspond to the classical conserved AID residues (11). Consensus sequences are also displayed as a function of channel type. Boxed residues correspond to sequence homology for L-type and non-L-type channels. Note that AID<sub>C</sub> has a sequence identical to that of AID<sub>D</sub>, and is thereafter referred to as AID<sub>C/D</sub>. B, Coomassie Blue-stained 9–15% SDS-polyacrylamide gel illustrating nonpurified control GST and various GST-I-II<sub>A</sub> fusion proteins containing the chimera AID sequences. These proteins result from the incubation of 200 µl of crude BL21 bacterial lysates with 40 µl of glutathione-agarose beads. The arrow denotes the position of the full-length GST-I-II<sub>A</sub> fusion proteins. C, autoradiogram demonstrating the specific binding of *in vitro* translated ³⁵S-β₄ binding to saturating concentrations of the chimeric GST fusion proteins (100 µl of crude bacterial lysates incubated with 40 µl of glutathione-agarose beads). The unbound ³⁵S-β₄ (translation), shown in the left band for comparison, illustrates that chimeric sequences have a maximum binding capacity. The molecular mass of ³⁵S-β₄ is 58 kDa.

differences are found in AID<sub>S</sub>, AID<sub>C</sub>, and AID<sub>D</sub>. More importantly, there are some significant differences in the distribution of charged residues between the two subfamilies of Ca<sub>v</sub> chan-
nels. Arg-5 of AID<sub>A</sub> is replaced by a Glu residue in L-type AID sequences, whereas Asn-8 and Lys-17 are replaced by Arg/Lys and Gln residues, respectively.

Replacing AID<sub>A</sub> from Ca<sub>v</sub>2.1 by an AID Sequence from An-
other Channel Does Not Inhibit Its Expression or Ability to Bind
the β<sub>4</sub> Subunit—To better understand the role of these AID
residues in calcium channel inactivation and β-dependent reg-
ulation, we made a set of chimeras in which we replaced the
AID<sub>A</sub> sequence from Ca<sub>v</sub>2.1 (20) by equivalent AID sequences
from Ca<sub>v</sub>1.1, Ca<sub>v</sub>1.2, or Ca<sub>v</sub>1.3 (L-type AID sequences) and from
Ca<sub>v</sub>2.2 (non-L-type AID sequences). All chimeric channels were
functionally expressed in Xenopus oocytes in the presence of
the β<sub>4</sub> subunit with average current densities ranging between
1.2 ± 0.2 and 13.4 ± 2.6 µA/µF (data not shown). We confirmed
that β<sub>4</sub> was indeed able to bind to the chimera channels by
directly testing the interaction of <sup>35</sup>S-β<sub>4</sub> subunit to various
crude preparations of chimeric GST-I-II<sub>A</sub> fusion proteins (Fig.
1B). The results illustrated in Fig. 1C confirm that β<sub>4</sub> is indeed
able to bind to the chimeric sequences. Affinity experiments
demonstrate that these interactions occur with an affinity
equivalent to that of the wild-type GST-I-II<sub>A</sub> linker (data not
shown). These data thus show that “nonconserved” AID resi-
dues are not essential to the binding of β subunits to the AID
sequences.

Replacing AID<sub>A</sub> from Ca<sub>v</sub>2.1 by an AID Sequence from an L-type Channel slows the Voltage-dependent Inactivation Rate—Next, we tested whether the chimeric channels differed in inactivation properties (Fig. 2). We compared the inactivation kinetics of wild-type Ca<sub>v</sub>2.1/β<sub>4</sub> channels with AID chimeric Ca<sub>v</sub>2.1/β<sub>4</sub> channels by measuring the time required for the current elicited at 20 mV to decrease to mid-amplitude. This paradigm was chosen because it allows a reliable comparison between the various channels expressed (see “Experimental Procedures”). The α<sub>2</sub>δ subunit was omitted from our expression experiments because it modifies inactivation in a β-dependent manner (22). The data obtained demonstrate that, in the presence of β<sub>4</sub>, there was no statistical difference in inactivation kinetics between wild-type Ca<sub>v</sub>2.1 and Ca<sub>v</sub>2.1 AID<sub>B</sub> channels, a result that is consistent with the close relatedness of their AID sequences. For a membrane potential of 20 mV, average HIT of Ca<sub>v</sub>2.1 is 152 ± 20 ms (n = 13) in this oocyte batch. In contrast, both Ca<sub>v</sub>2.1 AID<sub>S</sub> and Ca<sub>v</sub>2.1 AID<sub>C/D</sub> presented slower inactivation kinetics than wild-type Ca<sub>v</sub>2.1. The Ca<sub>v</sub>2.1 AID<sub>C/D</sub> chimera had the slowest inactivation kinetic with an average HIT value of 775 ± 78 ms (n = 13; slowing factor of 5.1 ± 1.1-fold). These data clearly demonstrate that the AID sequence contributes to the inactivation behavior of Ca<sub>v</sub>2.1 and are in agreement with the observation generally made that L-type channels have slower inactivation kinetics than non-L-type channels. However, one should not interpret these data to mean that the AID sequence is the unique determinant underlying kinetic differences between these channel types.

Nonconserved AID<sub>A</sub> Residues That Are Responsible for the Kinetic Differences between Wild-type Ca<sub>v</sub>2.1 and Chimera Ca<sub>v</sub>2.1 AID<sub>C</sub>—We aimed at identifying the AID amino acids that are responsible for such a drastic kinetic difference between wild-type and chimeric Ca<sub>v</sub>2.1 channels. We focused on the AID<sub>C/D</sub> chimera because its inactivation kinetics was the most divergent from that of the wild-type Ca<sub>v</sub>2.1. Fig. 3A illustrates several Ca<sub>v</sub>2.1 mutants in which individual nonconserved AID<sub>A</sub> residues were replaced by AID<sub>C</sub> residues located at equivalent positions. All these mutated AID sequences were able to interact with high affinity with the β<sub>4</sub> subunit (data not shown). Coexpression experiments of single mutated Ca<sub>v</sub>2.1 channels with β<sub>4</sub> reveal that, except for N390K, most nonconserved residues contribute to the slowing of inactivation ob-

a

<figure><img src="image_5.png" /></figure>

b

<figure><img src="image_6.png" /></figure>

FIG. 2. AID chimeras modify the inactivation kinetics of the Ca~v~2.1 channel. a, superimposed current traces elicited by depolarization to 20 mV for representative wild-type Ca~v~2.1 and Ca~v~2.1 AID~B~ (non-L-type AID sequences, left) and for Ca~v~2.1 AID~S~ and Ca~v~2.1 AID~C/D~ (L-type AID sequences, right). Wild-type Ca~v~2.1 current is shown for comparison in gray. All the channels were coexpressed with the β~4~ subunit. The bar for amplitude scaling corresponds to 0.4 µA (Ca~v~2.1 AID~A~, 0.25 µA (Ca~v~2.1 AID~B~, 0.2 µA (Ca~v~2.1 AID~S~, and 0.15 µA (Ca~v~2.1 AID~C/D~). b, average time required to reach half-inactivation for wild-type and chimera Ca~v~2.1 channels in the presence of β~4~. In parentheses are the numbers of cells used for the plot. For Ca~v~2.1 AID~S~ and Ca~v~2.1 AID~C/D~, the values were significantly slower than those of wild-type Ca~v~2.1 and chimera Ca~v~2.1 AID~B~ channels.

served with the Ca~v~2.1 AID~C/D~ chimera. The most striking
effect was observed with the R387E mutation with an average
HIT value of 660 ± 53 ms (n = 12). Very similar observations
have been made for this residue by various research groups (25,
26). However, none of these studies extended their investiga-
tion to closely positioned AID residues. Although several ami-
no-terminal residues slowed inactivation, the results obtained
with the AID~C/D~ chimera (Fig. 2) suggest that, in this region,
the effects of the individual residues are not additive as the
HIT value of the R387E mutation approached that of the
Ca~v~2.1 AID~C/D~ chimera. Overall, these data demonstrate that
more than one residue contributes to differences in inactivation
kinetics between AID~A~ and AID~C/D~. It was therefore of interest
to investigate whether the contribution of these amino acids to
inactivation was an intrinsic property of these residues them-
selves or whether their contribution was regulated by β~4~ sub-
unit association.

Two Individual Mutants (R387E and E388D) Affect the Sensitivity of the Channel to β Subunit Modulation—It is likely that the contribution of several AID residues to inactivation can be affected by the interaction of AID with the β₄ subunit.
<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

FIG. 3. Nonconserved AID residues implicated in inactivation in the presence of $\beta_4$ subunit. A, sequence alignment of AID$_A$ with AID$_C$ sequence. Single point mutations from AID$_A$ to AID$_C$ residues are also shown. B, half-inactivation time of Ca$_{2}$I AID$_A$ mutants in the presence of $\beta_4$. Representative current traces are shown at a test potential of +20 mV for wild-type (wt) Ca$_{2}$I and Ca$_{2}$I R387E (left). Average kinetic behaviors are also shown (right). Amplitude bar scale, 0.4 µA (wt Ca$_{2}$I) and 0.11 µA (Ca$_{2}$I R387E). The number of oocytes studied are shown in parentheses.

To address this point, we also analyzed the effects on kinetics of individual AID mutations in the absence of $\beta_4$ subunit (Fig. 4A). HIT values for wild-type Ca$_{2}$I channels were quite variable at 20 mV and may reflect differences in the conformational states of the channel (see “Discussion”). The average HIT value is 79 $\pm$ 13 ms ($n$ = 11) in this batch of oocytes. Channels with individual AID mutated residues were all functional in the absence of $\beta_4$ subunit. The HIT value was reduced by a factor of ~2-fold for three channel mutants: Ca$_{2}$I R387E (31 $\pm$ 7 ms, $n$ = 6), Ca$_{2}$I E388D (34 $\pm$ 6 ms, $n$ = 9), and Ca$_{2}$I N390K (44 $\pm$ 4 ms, $n$ = 8). In contrast to R387E, E388D, and N390K mutations, the average HIT value was increased by almost a factor of 2 for S397T (105 $\pm$ 11 ms, $n$ = 6) and K398Q (127 $\pm$ 13 ms, $n$ = 14) mutants (Fig. 4A). Curiously, however, some of the mutant channels that displayed the slowest inactivation kinetics in the presence of $\beta_4$ subunit were also those that had among the fastest inactivation kinetics in the absence of $\beta_4$ subunit (R387E and E388D). Hence, these data suggest that the contribution of several AID amino acids to inactivation can be modulated by the $\beta$ subunit. A plot of the $\beta_4$-induced slowing factor for wild-type Ca$_{2}$I, and for each of the mutant channels, illustrates that all channels have slower inactivation kinetics in the presence of $\beta_4$ subunit (Fig. 4B). Most mutant channels behave almost similarly to the wild-type channel (slowing factors ranging from 2.4 $\pm$ 0.4-fold for Ca$_{2}$I to 5.4 $\pm$ 0.6-fold for Ca$_{2}$I E394D), suggesting that they have an intrinsic ability to regulate voltage-dependent inactivation. In contrast, Ca$_{2}$I R387E and Ca$_{2}$I E388D, with slowing factors of 21.3 $\pm$ 4.3- and 11.0 $\pm$ 1.4-fold, respectively, are clearly different. Remarkably, one should note that the Ca$_{2}$I E388D mutant channel differs from wild-type Ca$_{2}$I by only a methylene group in the side chain of the residue. These data illustrate that nonconserved AID residues, particularly those lo-

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

FIG. 4. Nonconserved AID residues implicated in inactivation in the absence of $\beta_4$ subunit and effect of the mutation on $\beta$ regulation. A, half-inactivation time for Ca$_{2}$I AID$_A$ mutants in the absence of $\beta_4$ subunit. Representative current traces are shown at a test potential of +20 mV for wild-type (wt) Ca$_{2}$I, Ca$_{2}$I E388D, and Ca$_{2}$I K398Q (left). Average kinetic behaviors are also displayed (right). Amplitude bar scale, 0.1 $\mu$A (wild-type Ca$_{2}$I), 0.15 $\mu$A (Ca$_{2}$I K398Q), and 0.08 $\mu$A (Ca$_{2}$I E388D). B, the $\beta_4$-induced slowing in inactivation kinetics was studied by the score of the average ratio between half-inactivation values for each channel in the presence and in the absence of $\beta_4$. The upper panel illustrates the difference in kinetic slowing induced by $\beta_4$ subunit for wild-type Ca$_{2}$I and Ca$_{2}$I E388D. Amplitude bar scale, 0.1 $\mu$A (wild-type Ca$_{2}$I), 0.7 $\mu$A (wild-type Ca$_{2}$I/β$_4$), 0.08 $\mu$A (Ca$_{2}$I E388D), and 0.5 $\mu$A (Ca$_{2}$I E388D/β$_4$). In both A and B, the number of oocytes studied is shown in parentheses.

cated at the amino terminus of AID, contribute significantly to the kinetic behavior of the channel. Mutation of two of these residues affect the sensitivity of the channel to the $\beta$ subunit modulation. Next, we examined the effect of these AID amino acid substitutions on the steady-state inactivation of Ca$_{2}$I and tested whether similar predictions could be made with regard to $\beta$ subunit regulation.

Arg-387 and Glu-388 Are Also Two Critical Residues for the Regulation of Steady-state Inactivation by the $\beta_4$ Subunit—Besides differing by their inactivation kinetics, there is also evidence that the steady-state inactivation of L- and non-L-type channels is differently regulated by $\beta$ subunits. The steady-state inactivation of non-L-type channels is extensively modified by $\beta$ subunits, whereas that of L-type channels is more discretely altered. An initial investigation of the steady-state inactivation properties of the chimera channels, in the presence of the $\beta_4$ subunit, reveal differences in half-inactivation potential (HIP) values between L-type and non-L-type Ca$_{2}$I chimeras. For instance, the average HIP values observed with Ca$_{2}$I AID$_S$ and Ca$_{2}$I AID$_{C/D}$ were -28.7 $\pm$ 7.9 mV ($n$ = 6) and
<figure><img src="image_11.png" /><figcaption>FIG. 5. Effect of individual AID mutations on steady-state voltage-dependent inactivation and β₄ dependence. A, average HIP values for wild-type and mutant Caₓ₂.₁ channels in the absence (filled symbol) and presence (open symbol) of β₄ subunit. Average shift values and direction of shift are shown by arrows for each channel (mean ± S.E.). The number of oocytes studied are shown in parentheses. B, average half-activation potentials for wild-type and mutant Caₓ₂.₁ channels in the absence (black bars) and presence (gray bars) of β₄ subunit (mean ± S.E.). The number of oocytes studied are in parentheses, whereas the average current density at 20 mV are shown in brackets (in µA/µF). For comparison, the average current density for noninjected oocytes is 0.02 µA/µF (n = 8).</figcaption></figure>

-26.8 ± 2.3 mV (n = 7), thus more depolarized than the HIP value of -36.4 ± 6.7 mV (n = 11) for the wild-type Caₓ₂.₁ channels (data not shown). These data thus indicate that AID also plays a role in the control of steady-state inactivation, which prompted us to test the ability of the β₄ subunit to shift the voltage dependence of inactivation of the mutant Caₓ₂.₁ channels. Several conclusions can be drawn from this study. First, the formal HIP values are not identical among the mutants, both in the absence and presence of β₄ subunit, indicating that several residues may play a function in the voltage dependence of inactivation (Fig. 5A). This observation is in agreement with the observation made on the rate of inactivation, again suggesting that several AID residues contribute to the differences in inactivation behavior between wild-type Caₓ₂.₁ and chimera Caₓ₂.₁ AIDc/D. Second, the amplitude of β₄-induced shift is closely related for most residues. The shifts observed were mostly toward hyperpolarized values and ranged between -12.9 mV (M393L) and -20.5 mV (N390K), with an average value of -16.2 mV for wild-type Caₓ₂.₁. Third, the HIP values observed for Caₓ₂.₁ R387E and Caₓ₂.₁ E388D in the absence of β₄ and their β-dependences were significantly different from that of wild-type Caₓ₂.₁. Caₓ₂.₁ R387E and Caₓ₂.₁ E388D have average HIP values of -38.0 ± 2.5 mV (n =

7) and -40.1 ± 2.4 mV (n = 8), respectively. Coexpression of β₄ results in a modification of the HIP value of Caₓ₂.₁ R387E only, which shifts toward depolarized values, and becomes -28.2 ± 1.2 mV (n = 14), whereas the HIP value of Caₓ₂.₁ E388D remains unchanged with an average value of -40.8 ± 2.0 mV (n = 8). Thus, there is a significant change in β-dependence for both R387E and E388D mutants, which confers to these mutated Caₓ₂.₁ a property that is shared by L-type channels only. It is also interesting to note that the HIP values of Caₓ₂.₁ R387E and Caₓ₂.₁ E388D in the absence of β₄ are in close agreement with the HIP value of Caₓ₂.₁ in the presence of β₄. These data suggest that these two mutations may mimic the effect of β₄ on steady-state channel inactivation of Caₓ₂.₁. Additional arguments in favor of this interpretation will further be developed in this report. Of note, the effects of these mutations on inactivation were not associated with defects in the activation process (Fig. 5B). By measuring peak currents, we found that half-activation potentials (HAP) were closely related in the absence of β₄ subunit (HAP values of 2.7 ± 1.2 mV (Caₓ₂.₁, n = 9), 6.3 ± 0.9 mV (Caₓ₂.₁ R387E, n = 7), and 3.2 ± 0.7 mV (Caₓ₂.₁ E388D, n = 9)). Similarly, in the presence of β₄, we observed average HAP values of -9.9 ± 0.5 mV (Caₓ₂.₁, n = 8), -9.7 ± 0.7 mV (Caₓ₂.₁ R387E, n = 8), and -8.5 ± 0.7 mV (Caₓ₂.₁ E388D, n = 9). For other mutants (i.e. I385L), HAP values and β-dependence were slightly different, indicating that their implication in the β-dependent shift of activation remains possible (14). These results demonstrate that there are two nonconserved residues in AID that are essential for the regulation of the kinetics and voltage dependence of inactivation by the β₄ subunit.

The I-II Loop of Caₓ₂.₁ Interacts with Several Other Cytoplasmic Loops of the Channel, All Potentially Contributing to the Process of Inactivation—Recently, several studies have postulated that the I-II loop may act as an inactivation particle by physically occluding the pore (7, 27, 28). Although this hypothesis may seem attractive, the receptor site for such an inactivation particle has not been identified. In a collaborative study, we have presented evidence that the I-II loop may interact with other sequences within the channel (27). We confirmed this finding by studying the interaction of *in vitro* translated ³⁵S-I-IIA loop with several Caₓ₂.₁ GST fusion proteins (Fig. 6). Various GST fusion proteins of the Caₓ₂.₁ channel have been illustrated in Fig. 6A. At fusion protein concentrations of 1 µM, we detect a clear interaction of ³⁵S-I-IIA with the amino terminus, the I-II loop itself (self-association), the III-IV loop, and the carboxyl terminus (Fig. 6B). In contrast, there is no interaction with control GST or GST-II-IIIA fusion proteins, confirming the specificity of these binding experiments. The binding affinity between the I-II loop and GST-III-IVA was 350 nM (data not shown). Fig. 6C summarizes these various detected intramolecular interactions of Caₓ₂.₁. Obviously, these findings do not provide any information on the number of interactions that the I-II loop is able to manage simultaneously. However, these results clearly indicate that the process of inactivation cannot be analyzed in a simplistic manner without considering the structural and functional consequences of sequence swapping within Caₓ₂.₁. We next determined which one of these interactions may be important to inactivation. This question was treated in the absence of the β₄ subunit because β association to the I-II loop may produce a structural reorganization in Caₓ₂.₁ interactions.

Overexpression of the III-IV Loop Modifies Caₓ₂.₁ Inactivation in a Manner Similar to R387E and E388D Mutations—We investigated the relative importance of the various I-II loop interactions in inactivation by attempting to competitively inhibit them by the overexpression of cytoplasmic Caₓ₂.₁ se-
<figure><img src="image_12.png" /></figure>

FIG. 6. The I-II<sub>A</sub> loop is able to interact with several cytoplasmic domains of Ca<sub>v</sub>2.1. A, schematic diagram illustrating the various GST-Ca<sub>v</sub>2.1 constructs that were used for the binding of the <sup>35</sup>S-I-II<sub>A</sub> loop. Similar CD8-Ca<sub>v</sub>2.1 constructs were used for coexpression with Ca<sub>v</sub>2.1 in Fig. 7 except that the carboxyl terminus of Ca<sub>v</sub>2.1 was split into two constructs (Ct1<sub>A</sub> and Ct2<sub>A</sub>). B, the full-length I-II<sub>A</sub> loop of Ca<sub>v</sub>2.1 was translated *in vitro* and labeled with [<sup>35</sup>S]methionine, and binding onto various GST fusion proteins was assayed. The <sup>35</sup>S-I-II<sub>A</sub> loop was found to interact with GST-Nt<sub>A</sub>, GST-I-II<sub>A</sub>, GST-III-IV<sub>A</sub>, and GST-Ct<sub>A</sub>. Fusion protein concentration was 1 µM. Overnight autoradiogram exposure time. Top insets illustrate the equivalent quantities of GST fusion proteins used. C, schematic diagram illustrating the various intramolecular interactions of the I-II loop. Note that the I-II<sub>A</sub> loop binds also onto itself, probably reflecting three-dimensional structural constraints.

quences. Various chimera CD8-Ca,2.1 constructs were used to facilitate cell surface detection and to anchor these Ca,2.1 sequences to the membrane. In these constructs, the cytoplasmic sequence of the αCD8 chain was replaced by various Ca,2.1 cytoplasmic sequences (also depicted in Fig. 6A). Membrane anchoring should also increase the probability of association of these sequences to Ca,2.1. According to metabolic labeling and immunoprecipitation experiments, the various CD8-Ca,2.1-Myc constructs all expressed well in *Xenopus* oocytes (Fig. 7A, top panel). All these constructs expressed with much better

<figure><img src="image_13.png" /></figure>

FIG. 7. Role of the III-IV loop in the inactivation of Ca<sub>v</sub>2.1 alone. A, effect of the various CD8-Ca<sub>v</sub>2.1-Myc constructs on the average HIT value of Ca<sub>v</sub>2.1. Mean ± S.E. and *n* = 127 oocytes. Top insets illustrate the expression level of the various <sup>35</sup>S-CD8-Ca<sub>v</sub>2.1-Myc constructs in oocytes (*n* = 30) following metabolic labeling with [<sup>35</sup>S]methionine and anti-Myc immunoprecipitation. The autoradiogram was exposed for 24 h. B, representative current traces at 20 mV for Ca<sub>v</sub>2.1 and Ca<sub>v</sub>2.1 coexpressed with CD8-III-IV<sub>A</sub> illustrating the increase in inactivation rate. Amplitude bar scale, 0.1 µA (Ca<sub>v</sub>2.1) and 0.08 µA (Ca<sub>v</sub>2.1 + CD8-III-IV<sub>2.1</sub>). C, average steady-state inactivation curves for Ca<sub>v</sub>2.1 alone (*n* = 8) or coinjected with either 10 µM III-IV<sub>A</sub> peptide (*n* = 7) or a control peptide (*n* = 4).

efficacies at the plasma membrane than the Ca<sub>v</sub>2.1 channel itself (data not shown), thereby preventing the possibility to specifically study the interaction of Ca<sub>v</sub>2.1 and the various CD8-Ca<sub>v</sub>2.1 proteins by biochemical techniques. We next tested the effect of expressing these constructs on the HIT values of Ca<sub>v</sub>2.1 (Fig. 7A). These experiments were performed on a single oocyte batch to avoid inter-batch variability. The data illustrate that the control construct, CD8-stop, has no effect on the average HIT value of Ca<sub>v</sub>2.1. A similar observation was made for several chimeric CD8 constructs, including CD8-I-II<sub>A</sub>. It should be noted that a lack of effect may simply imply that the expressed sequences were not able to disorganize the intramolecular structure of the channel. In contrast, we found that CD8-III-IV<sub>A</sub> significantly increases the rate of inactivation of Ca<sub>v</sub>2.1 (average HIT value of 18.1 ± 1.4 ms at 20 mV (*n* = 6) instead of 50.3 ± 5.4 ms (*n* = 16) for CD8-stop; Fig. 7B). We further confirmed that the decrease in HIT value produced by CD8-III-IV<sub>A</sub> was a result of the III-IV<sub>A</sub> sequence itself because co-injection of a 40-mer peptide derived from the III-IV<sub>A</sub> loop (Ser-1535 to Ser-1574, 10 µM) produced a similar HIT value of 20.6 ± 2.2 ms (*n* = 8; Fig. 7A) at 20 mV. Interestingly, the HIT values obtained in the presence of III-IV<sub>A</sub> are also reminiscent of the HIT values of Ca<sub>v</sub>2.1 R387E and Ca<sub>v</sub>2.1 E388D (Fig. 4A). Because of these similarities, we tested the possibility that the injection of 10 µM III-IV<sub>A</sub> peptide produces a shift in the steady-state inactivation of Ca<sub>v</sub>2.1 (Fig. 7C). The peptide produces a
A

<figure><img src="image_14.png" /></figure>

B

<figure><img src="image_15.png" /></figure>

C

<figure><img src="image_16.png" /></figure>

FIG. 8. Similarities in the mechanisms of action of R387E and E388D AID mutants and the III-IV<sub>A</sub> peptide. A, left, Coomassie Blue-stained gel illustrating purified GST and GST-III-IV<sub>A</sub> fusion proteins; right, autoradiogram illustrating the binding of various <sup>35</sup>S-I-II<sub>A</sub> mutant sequences onto 1 µM GST-III-IV<sub>A</sub>. The binding of <sup>35</sup>S-I-II<sub>A</sub> E388D and <sup>35</sup>S-I-II<sub>A</sub> R387E is reduced or abolished, respectively. B, lack of effect of 10 µM of III-IV<sub>A</sub> peptide on HIT values for Ca<sub>v</sub>2.1 E388D. Left, representative current traces at 20 mV; right, box plot representation of HIT values. C, average steady-state inactivation curves for Ca<sub>v</sub>2.1 E388D alone (n = 8) or coinjected with the III-IV<sub>A</sub> peptide (n = 8).

change of the HIP value from -20.2 ± 1.6 mV (control Ca<sub>v</sub>2.1, n = 8) to -37.1 ± 2.4 mV (III-IV<sub>A</sub> peptide, n = 7), whereas the injection of a nonrelevant peptide had no effect (average HIP of -22.5 ± 0.8 mV, n = 4). A similar shift in HIP value was observed with the CD8-III-IV<sub>A</sub> construct (HIP of -41.2 ± 2.5 mV, n = 6; data not shown). Thus, these data indicate that the interaction between the I-II loop and the III-IV loop contributes to the process of inactivation in the absence of the β₄ subunit. The striking similarities observed between the inactivation properties of Ca<sub>v</sub>2.1 R387E, Ca<sub>v</sub>2.1 E388D, and Ca<sub>v</sub>2.1 coexpressed with the III-IV<sub>A</sub> sequence suggest that both R387E and E388D may regulate inactivation by selectively disrupting the interaction between the I-II and III-IV loops.

Like the III-IV Peptide, R387E and E388D Mutations Also Weaken the Functional I-II/III-IV Interaction—Fig. 8A illus-

trates that the binding of <sup>35</sup>S-I-II<sub>A</sub> R387E or <sup>35</sup>S-I-II<sub>A</sub> E388D onto 1 µM GST-III-IV<sub>A</sub> is significantly reduced compared with that of wild-type <sup>35</sup>S-I-II<sub>A</sub>. A weak interaction seems to persist for the E388D mutant, suggesting that the affinity of binding is reduced. Importantly, other I-II loop mutants, which do not alter β regulation of Ca<sub>v</sub>2.1 inactivation, keep normal interaction levels with GST-III-IV<sub>A</sub>. To further functionally confirm that the AID mutants act by disrupting the normal I-II/III-IV interaction, we tested the effect of the III-IV<sub>A</sub> peptide on the kinetics and voltage dependence of inactivation of Ca<sub>v</sub>2.1 E388D (Fig. 8, B and C). HIT values observed at 20 mV for Ca<sub>v</sub>2.1 E388D were 34.1 ± 5.6 and 38.3 ± 1.3 ms in the absence and presence of the peptide, respectively (Fig. 8B). In addition, HIP values were -40.1 ± 2.4 mV (n = 8, Ca<sub>v</sub>2.1 E388D) and -38.7 ± 1.7 mV (n = 8, Ca<sub>v</sub>2.1 E388D + peptide III-IV<sub>A</sub>). The lack of effect of the peptide on the Ca<sub>v</sub>2.1 E388D strongly suggests that both the mutation and the peptide act by a similar mechanism to alter inactivation of the channel. This mechanism appears to be the disruption of the interaction between the I-II loop and the III-IV loop. Interestingly, the observation that Arg-387 and Glu-388 are the two only nonconserved AID residues, the contribution of which to inactivation is more drastically modified by β subunit association, leads to the hypothesis that β₄ may also modify the interaction between the I-II loop and the III-IV loop. In favor of such an interpretation is the finding that β₄ subunit produces a shift in the HIP value of Ca<sub>v</sub>2.1 similar to that observed with R387E, E388D or the III-IV<sub>A</sub> peptide. We therefore tested the ability of the III-IV peptide to modify the inactivation of Ca<sub>v</sub>2.1/β₄ channels.

The III-IV Peptide Is Not Active on Ca<sub>v</sub>2.1 Inactivation in the Presence of the β₄ Subunit—Fig. 9A illustrates that overexpression of CD8-III-IV<sub>A</sub>, or injection of the III-IV<sub>A</sub> peptide, has no effect on the HIT value of Ca<sub>v</sub>2.1/β₄, thereby clearly demonstrating that the binding of β₄ onto the I-II loop prevents the effect of exogenous III-IV<sub>A</sub> peptide on the channel. This observation further rules out the possibility that the III-IV<sub>A</sub> peptide acts on inactivation by binding onto another receptor structure than the I-II loop. Our results are thus consistent with the interpretation that R387E or E388D, excess of the III-IV loop, or the β₄ subunit all act on inactivation by a common step, which is the disruption of the interaction between the I-II loop and the III-IV loop.

DISCUSSION

In this report, we present compelling evidence that several nonconserved AID residues are important in controlling inactivation in voltage-dependent calcium channels. Mutations of the AID<sub>A</sub> sequence toward residues found in AID<sub>C</sub> generally produced a slowing of the inactivation rate of Ca<sub>v</sub>2.1. Our observations thus extend previous findings on the contribution of Arg-387 to inactivation in non-L-type channels (25, 26). We also demonstrate that mutation of two of the nonconserved residues (Arg-387 and Glu-388) modify the extent to which β subunit regulates inactivation. This is an essential finding because it may explain some of the differences observed in the regulation of inactivation by β subunits between L- and non-L-type channels (14). Finally, we demonstrate for the first time that the III-IV loop contributes to inactivation through an interaction with the I-II loop. This intramolecular interaction can be disrupted in three different ways: (i) mutation of Arg-387 or Glu-388 residues, (ii) overexpression of the III-IV loop sequence or injection of an excess III-IV peptide, and (iii) coexpression of the β₄ subunit. Loss of the native interaction between the I-II loop and the III-IV loop inhibits the contribution of the III-IV loop to Ca<sub>v</sub>2.1 inactivation.

In the following sections, we will discuss how our data can be integrated in current models of calcium channel inactivation.
A

<figure><img src="image_17.png" /></figure>

B

<figure><img src="image_18.png" /></figure>

FIG. 9. In the presence of $β_4$ subunit, the III-IV peptide no longer regulates the inactivation of $Ca_v$2.1. A, lack of effect of 10 µM III-IV$_A$ peptide on HIT values for $Ca_v$2.1 E388D coexpressed with $β_4$ subunit. Left, representative current traces at 20 mV; right, box plot representation of HIT values. B, hypothetical model calcium channel reorganization by the I-II loop mutation, the III-IV peptide, and the $β$ subunit. Inactivation is altered in various ways because of the crucial role of the I-II loop in this process.

We also illustrate how $β$ subunits may be implicated in inactivation by diverting some of the I-II determinants.

*Multiplicity of Determinants Implicated in Inactivation*—In calcium channels, the multiplicity of molecular determinants implicated in inactivation is impressive. Despite this apparent complexity, one can nevertheless classify these determinants into two types: transmembrane segments and cytoplasmic elements. Similar to potassium channels (29) and possibly sodium channels, several lines of evidence point to the importance of pore-lining S6 segments in calcium channel inactivation. Sequence swapping illustrates the essential contribution of the IS6 (3), IIS6 (7), IIIS6 (30), and IVS6 (31) segments to fast inactivation. Additional elements have however also been identified in cytoplasmic sequences, the amino terminus (13), the I-II loop (4, 26), and the carboxyl terminus (5). For instance, a Val insertion, 18 residues carboxyl to AID, produces a slowing of inactivation kinetics and affects the steady-state inactivation (4). Recently, it was proposed that the domain II and III S6 segments and the I-II linker work in concert to set the rate of calcium channel inactivation (7). Our own finding that the III-IV loop contributes to voltage-dependent inactivation in a $β$-dependent manner further extends the list of structural de-

terminants implicated in this process. Obviously, the interpretation of the respective role of each one of these determinants in inactivation requires a clearer representation of what the inactivated state of a calcium channel might be.

*Models of the Inactivated State of Voltage-dependent Calcium Channels*—Two noncontradictory models have been proposed that may explain how inactivation proceeds in voltage-dependent calcium channels (for review, see Stotz and Zamponi (Ref. 28)).

It was recently proposed by Stotz and co-workers (7) that the I-II linker might act as an inactivation particle that would physically occlude the pore of the channel by docking in part to the S6 segments of domains II and III. This model is thus analogous to the ball and chain, and to the hinged-lid mechanisms described in $K^+$ and $Na^+$ channels, respectively (29, 32). Several observation would favor this model. First, it was found that overexpression of the I-II linker increases the inactivation rate of $Ca_v$2.1 (9), although, in this case, the effect could also be related to an inhibition of $β$ subunit association. Second, it was found that G proteins, known to bind onto the I-II loop of several $Ca_v$ channels, may affect ion channel selectivity (33). Third, it was demonstrated that some charged piperidine mol-
ecules can bind to both the pore structure and to the I-II loop. This hypothesis remains also attractive because it would represent an unifying mechanism whereby inactivation would occur through pore occlusion in voltage-gated cation channels.

The crystal structure of a potassium channel has provided interesting clues on the role of S6 segments in channel gating (34) and for an alternative model of inactivation. According to this crystal structure, the various S6 segments would be positioned as an inverted teepee. It is thus conceivable that inactivation could proceed through a pore collapse mediated by a repositioning of various S6 segments. Intramembrane movement of the S6 segment is expected to explain, for instance, the use dependence of phenylalkylamine block (6). This model would thus be similar to the C-type inactivation described for $K^+$ channels in which inactivation is thought to occur through S6-mediated pore constriction (29). Data from Spaetgens and Zamponi (8) further support the notion that all four transmembrane domains of $Ca_{v}2.3$ contribute to fast inactivation. Evidence that the pore-forming S6 segments are packed against each other stems from the observation that residues of the IIS6 segment can simultaneously affect inactivation and phenylalkylamine sensitivity, a drug that binds onto the IVS6 segment (35). In this model, the contribution of cytoplasmic sequences to inactivation would not occur through a hinged-lid mechanism, but via the modulation of the relative mobility and positioning of the various S6 segments. The observation that $\beta$ subunits, syntaxin, and calmodulin, all regulate inactivation by binding, respectively, onto the I-II linker (11), the II-III loop (36), and the carboxyl terminus (37) is also in line with this hypothesis.

It should be emphasized that these two models are not necessarily contradictory because inactivation may represent the sum of multiple mechanisms, as in the case of *Shaker* potassium channels, in which a complex interplay between N- and C-type inactivation is demonstrated (29).

*How Do Our Data Fit with These Two Models?*—Our data provide novel evidence for the implication of the I-II loop in inactivation. We found that the I-II loop binds onto multiple cytoplasmic domains; of which several were previously identified as critical determinants implicated in inactivation. These include the amino (12, 13) and the carboxyl termini (5, 16, 37). In fact, this close structural link between these various structural parts of $Ca_{v}2.1$ channel was suspected for several reasons. For instance, the $\beta_{4}$ subunit, which anchors onto the I-II loop, does also associate to the amino and the carboxyl terminus (14). Similarly, the $G\beta\gamma$ complex also appears capable to bind to the I-II loop (38), the amino terminus (39), and the carboxyl terminus (40) further suggesting a structural proximity between these various $Ca_{v}$ sequences.

The multiple possibilities of interaction of the I-II loop with the various cytoplasmic sequences of $Ca_{v}2.1$ has several structural and functional implications. First, at least one of these interactions is involved in inactivation. We found that overexpression of the III-IV loop facilitates $Ca_{v}2.1$ inactivation, presumably by competitively freeing the native I-II/III-IV interaction. A similar effect on inactivation is evidenced following mutation of residues at position 387 and 388, mutations that also weakened the interaction between I-II and III-IV. Interestingly, the observation that overexpression of a free-moving I-II loop modifies inactivation (9) can now also be interpreted as a modification in the balance of intramolecular interactions. Second, although further complicated biochemical analyses would be required to validate this hypothesis, it is unlikely that all these interactions may occur simultaneously in the channel. We found previously that $Ca_{v}2.1$ inactivates along three components, two inactivating and one noninactivating (41). One

could therefore assume that the various modes of $Ca_{v}2.1$ inactivation are supported by different sets of intramolecular interactions in which the I-II loop is implicated. Although a more careful kinetic examination would be required, the variability in HIT values, which we observe for $Ca_{v}2.1$ channels expressed alone, is consistent with the concept of a variability in intramolecular interactions between cytoplasmic sequences.

These findings seem to fit nicely with the two models of fast calcium channel inactivation. On one hand, binding of the I-II loop to several other cytoplasmic segments may restrict the mobility of the I-II loop and the ease at which it may occlude the ionic pore. In that respect, excess III-IV loop or mutation of Arg-387 or Glu-388 would facilitate the mobility of the I-II loop and thereby explain a faster rate of inactivation. Additionally, we observed less kinetic variability for $Ca_{v}2.1$ channels coexpressed with the III-IV loop (Fig. 7), a result that is again consistent with a simplification in the internal set of molecular interactions. On the other hand, the various intramolecular interactions of $Ca_{v}2.1$ probably regulate inactivation by altering the relative positioning of the various S6 segments. In that context, a disruption in the interaction between I-II and III-IV loop would also favor an easier and faster pore constriction by the COOH-terminal extremities of the S6 segments.

Among the two potential models of inactivation, we favor the second one. First, we have no strong evidence to believe that the various cytoplasmic sequences, on which the I-II loop binds, may represent part of the receptor site for the I-II loop in a hinged-lid mechanism of inactivation. In fact, the experimental evidence points to the opposite. By eliminating the III-IV loop as a potential receptor site of the I-II loop, we actually facilitate inactivation rather than inhibiting it. Additionally, overexpression of the other loops has no effect on inactivation, suggesting that these sequences are not required for fast inactivation by the I-II loop. Second, it is difficult to understand why the lid efficiency of the I-II loop is not affected by the binding of the $G\beta\gamma$ complex. The data presented here and elsewhere point rather to an immobilization of this loop through intramolecular interactions (27), $\beta$ subunit association (11), and G protein regulation (26, 38). However, further experimental testing will be required to evaluate these various models of inactivation.

In contrast to the I-II loop, there is no reason to believe that the III-IV loop has a central function in calcium channel inactivation. First, the III-IV loop is unable to affect the inactivation rate in the presence of $\beta$ subunit, suggesting that it does not behave as an inactivation moiety per itself. Second, binding of the $\beta$ subunit onto the I-II loop presumably releases the III-IV loop, which should therefore be free to occlude the pore more easily in the absence of conformational constraints: something that is not observed. Third, the kinetic effects of overexpressing the III-IV loop on $Ca_{v}2.1$ currents were strictly identical both in its peptide form (free-moving structure) or in its membrane bound form (plasma membrane-linked sequence), suggesting that conformational constrains did not hinder the peptide motif to reach its target binding site. Besides, pore blockers are expected to have molecular masses below 900 kDa and to be positively charged (42). Hence, our data strongly suggest that, unlike voltage-dependent sodium channels, the III-IV loop is not involved in inactivation according to a “hinged-lid” mechanism (32).

*How Do $\beta$ Subunits Take Over the Control of Inactivation?*—Changes in inactivation by $\beta$ subunits could proceed in several ways. First, $\beta$ subunit may reposition the I-II loop with regard to other essential structural elements. Such a conformational repositioning of the I-II loop has been proposed following the observation that the $\beta_{2a}$ subunit alters the piperidine receptor site (43). These data are in line with our own results showing
that there are indeed some residues that contribute to inactivation in a β-dependent manner. Second, the binding of the β subunit may facilitate conformational changes in the channel by competitively inhibiting the interactions of the I-II loop with several other cytoplasmic domains of the Ca²⁺,2.1. This mechanism would be equivalent to the competitive freeing of the I-II loop that was obtained with an excess III-IV peptide. The observation that the β₄ subunit shifts the voltage dependence of inactivation to values very similar to that obtained with the III-IV loop or Arg-387 and Glu-388 mutations is consistent with that concept. Preliminary experimental evidence suggest that intramolecular binding site disruption is indeed one of the mechanism of regulation adopted by the β₃ subunit (data not shown). Third, binding of the β subunit onto the I-II linker may slow down inactivation by altering its conformation or decreasing its mobility. We found, however, that the various β subunits kept their ability to distinctly regulate inactivation (data not shown), suggesting that conformational constraints, such as membrane attachment by palmitoylation in the case of β₂α or channel attachment by secondary sites in the case of β₄, play an essential role in secondary conformational constraints.

These various mechanisms of regulation by β subunits may all contribute to inactivation in voltage-dependent calcium channels and emphasize the need for an accelerated structural inspection of these molecules.

**Concluding Remarks**—In conclusion, our findings will facilitate the understanding of the role of β subunits in calcium channel voltage-dependent inactivation. In this report, we have addressed the mechanistic aspects of Ca²⁺,2.1 inactivation, and further analysis will be required to understand the physiological implication of each of the structural determinants identified so far. The implication of the III-IV loop in the inactivation process of native P/Q calcium channels that contain all their auxiliary subunits is dubious. However, the fact that a peptide derived from this loop is able to regulate channels that lack their β subunit but not those that are associated with β will represent a unique discriminative tool to segregate between native calcium channels with or without associated β subunit.

**Acknowledgment**—We are indebted to Dr. Kevin Campbell for critical reading of the manuscript.

REFERENCES

1. Forsythe, I. D., Tsujimoto, T., Barnes-Davies, M., Cuttle, M. F., and Takahashi, T. (1998) *Neuron* **20**, 797–807

2. Patil, P. G., Brody, D. L., and Yue, D. T. (1998) *Neuron* **20**, 1027–1038

3. Zhang, J.-F., Ellinor, P. T., Aldrich, R. W., and Tsien, R. W. (1994) *Nature* **372**, 97–100

4. Bourinet, E., Soong, T. W., Sutton, K., Slaymaker, S., Mathews, E., Monteil, A., Zamponi, G. W., Nargeot, J., and Snutch, T. P. (1999) *Nat. Neurosci.* **2**, 407–415

5. Soldatov, N. M., Zühlke, R. D., Bouron, A., and Reuter, H. (1997) *J. Biol. Chem.* **272**, 3560–3566

6. Hering, S., Berjukow, S., Aczel, S, and Timin, E. N. (1998) *Trends Pharmacol. Sci.* **19**, 439–443

7. Stotz, S. C., Hamid, J., Spaetgens, R. L., Jarvis, S. E., and Zamponi, G. W. (2000) *J. Biol. Chem.* **275**, 24575–24582

8. Spaetgens, R. L., and Zamponi, G. W. (1999) *J. Biol. Chem.* **274**, 22428–22436

9. Cens, T., Reste, S., Galas, S., and Charnet, P. (1999) *J. Biol. Chem.* **274**, 5483–5490

10. Wei, S.-K., Colecraft, H. M., DeMaria, C. D., Peterson, B. Z., Zhang, R., Kohout, T. A., Rogers, T. B., and Yue, D. T. (1999) *Circ. Res.* **86**, 175–184

11. Pragnell, M., De Waard, M., Mori, Y., Tanabe, T., Snutch, T. P., and Campbell, K. P. (1994) *Nature* **368**, 67–70

12. Walker, D., Bichet, D., Geib, S., Mori, E., Cornet, V., Snutch, T. P., Mori, Y., and De Waard, M. (1999) *J. Biol. Chem.* **274**, 12383–12390

13. Stephens, G. J., Page, K. M., Bogdanov, Y., and Dolphin, A. C. (2000) *J. Physiol.* **525**, 377–390

14. Walker, D., and De Waard, M. (1998) *Trends Neurosci.* **21**, 148–154

15. Yamaguchi, H., Hara, M., Strobeck, M., Fukasawa, K., Schwartz, A., and Varadi, G. (1998) *J. Biol. Chem.* **273**, 19348–19356

16. Walker, D., Bichet, D., Campbell, K. P., and De Waard, M. (1998) *J. Biol. Chem.* **273**, 2361–2367

17. Bichet, D., Cornet, V., Geib, S., Carlier, E., Volsen, S., Hoshi, T., Mori, Y., and De Waard, M. (2000) *Neuron* **25**, 177–190

18. De Waard, M., Pragnell, M., and Campbell, K. P. (1994) *Neuron* **13**, 495–503

19. Qin, N., Olcese, R., Zhou, J., Cabello, O. A., Birnbaumer, L., and Stefani, E. (1996) *Am. J. Physiol.* **271**, C1539–C1545

20. Mori, Y., Friedrich, T., Kim, M.-S., Mikami, A., Nakai, J., Ruth, P., Bosse, E., Hofmann, F., Flickerzi, V., Furuichi, T., Mikoshiba, K., Imoto, K., Tanabe, T., and Numa, S. (1991) *Nature* **350**, 398–402

21. Eppig, J. J., and Dumont, J. N. (1976) *In Vitro* **12**, 418–427

22. De Waard, M., Witcher, D. R., Pragnell, M., Liu, H., and Campbell, K. P. (1995) *J. Biol. Chem.* **270**, 12056–12064

23. Stea, A., Dubel, S. J., Pragnell, M., Leonard, J. P., Campbell, K. P., and Snutch, T. P. (1993) *Neuropharmacology* **32**, 1103–1116

24. Tomlinson, W. J., Stea, A., Bourinet, E., Charnet, P., Nargeot, J., and Snutch, T. P. (1993) *Neuropharmacology* **32**, 1117–1126

25. Berrou, L., Bernatchez, G., and Parent, L. (2001) *Biophys. J.* **80**, 215–228

26. Herlitze, S., Hockerman, G. H., Scheuer, T., and Catterall, W. A. (1997) *Proc. Natl. Acad. Sci. U. S. A.* **94**, 1512–1516

27. Restituito, S., Cens, T., Barrere, C., Geib, S., Galas, S., De Waard, M., and Charnet, P. (2000) *J. Neurosci.* **20**, 9046–9052

28. Stotz, S. C., and Zamponi, G. W. (2001) *Trends Neurosci.* **24**, 176–181

29. Hoshi, T., Zagotta, W. N., and Aldrich, R. W. (1991) *Neuron* **7**, 547–556

30. Tang, S., Yatani, A., Bahinski, A., Mori, Y., and Schwartz, A. (1993) *Neuron* **11**, 1013–1021

31. Döring, F., Degtiar, V. E., Grabner, M., Striessnig, J., Hering, S., and Glossmann, H. (1996) *J. Biol. Chem.* **271**, 11745–11749

32. Eaholtz, G., Scheuer, T., and Catterall, W. A. (1994) *Neuron* **12**, 1041–1048

33. Kuo, C. C., and Bean, B. P. (1993) *Nature* **365**, 258–262

34. Doyle, D. A., Cabral, J. M., Pfuetzner, R. A., Kuo, A., Gulbis, J. M., Cohen, S. L., Chait, B. T., and McKinnon, R. (1998) *Science* **280**, 69–77

35. Hering, S., Aczél, S., Kraus, R. L., Berjukow, S., Striessnig, J., and Timin, E. N. (1997) *Proc. Natl. Acad. Sci. U. S. A.* **94**, 13323–13328

36. Bezprozvanny, I., Scheller, R. H., and Tsien, R. W. (1995) *Nature* **378**, 623–626

37. Lee, A., Wong, S. T., Gallagher, D., Li, B., Storm, D. R., Scheuer, T., and Catterall, W. A. (1999) *Nature* **399**, 155–159

38. De Waard, M., Liu, H., Walker, D., Scott, V. E. S., Gurnett, C. A., and Campbell, K. P. (1997) *Nature* **385**, 446–450

39. Canti, C., Page, K. M., Stephens, G. J., and Dolphin, A. C. (1999) *J. Neurosci.* **19**, 6855–6864

40. Qin, N., Platano, D., Olcese, R., Stefani, E., and Birnbaumer, L. (1997) *Proc. Natl. Acad. Sci. U. S. A.* **94**, 8866–8871

41. De Waard, M., and De Waard, M. (1995) *J. Physiol.* **485**, 619–634

42. Hille, B. (1992) *Ionic Channels of Excitable Membranes*, 2nd Ed., Sinauer Associates, Inc., Sunderland, MA

43. Zamponi, G. W., Soong, T. W., Bourinet, E., and Snutch, T. P. (1996) *J. Neurosci.* **16**, 2430–2443