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Identification of Glycosylation Sites Essential for Surface Expression of the CaVα2δ1 Subunit and Modulation of the Cardiac CaV1.2 Channel Activity*

Alteration in the L-type current density is one aspect of the electrical remodeling observed in patients suffering from cardiac arrhythmias. Changes in channel function could result from variations in the protein biogenesis, stability, post-translational modification, and/or trafficking in any of the regulatory subunits forming cardiac L-type Ca2+ channel complexes. CaVα2δ1 is potentially the most heavily N-glycosylated subunit in the cardiac L-type CaV1.2 channel complex. Here, we show that enzymatic removal of N-glycans produced a 50-kDa shift in the mobility of cardiac and recombinant CaVα2δ1 proteins. This change was also observed upon simultaneous mutation of the 16 Asn sites. Nonetheless, the mutation of only 6/16 sites was sufficient to significantly 1) reduce the steady-state cell surface fluorescence of CaVα2δ1 as characterized by two-color flow cytometry assays and confocal imaging; 2) decrease protein stability estimated from cycloheximide chase assays; and 3) prevent the CaVα2δ1-mediated increase in the peak current density and voltage-dependent gating of CaV1.2. Reversing the N348Q and N812Q mutations in the non-operational sextuplet Asn mutant protein partially restored CaVα2δ1 function. Single mutation N663Q and double mutations N348Q/N468Q, N348Q/N812Q, and N468Q/N812Q decreased protein stability/synthesis and nearly abolished steady-state cell surface density of CaVα2δ1 as well as the CaVα2δ1-induced up-regulation of L-type currents. These results demonstrate that Asn-663 and to a lesser extent Asn-348, Asn-468, and Asn-812 contribute to protein stability/synthesis of CaVα2δ1, and furthermore that N-glycosylation of CaVα2δ1 is essential to produce functional L-type Ca2+ channels.

The regulation of Ca 2ϩ influx in cardiac cells is critical to the generation of the force necessary for the myocardium to meet the physiological needs of the body (1). In resting cells, intracellular free ionized Ca 2ϩ is maintained at a low concentration (high nanomolar range) by the concerted action of mechanisms that prevent Ca 2ϩ entry, promote its extrusion (mostly via the Na ϩ /Ca 2ϩ exchanger), and ensure its storage in the sarcoplasmic reticulum (2). Ca 2ϩ entry is mediated mainly by the cardiac L-type Ca 2ϩ channel, which is central to the initiation of excitation-contraction coupling via Ca 2ϩ -induced Ca 2ϩ release from the sarcoplasmic reticulum. Regulation of the L-type Ca 2ϩ current has profound physiological significance. Indeed, alterations in density or the activation/inactivation gating of L-type Ca 2ϩ channels have been implicated in a variety of cardiovascular diseases (3,4), including cardiac arrhythmias such as atrial fibrillation (5)(6)(7)(8), heart failure (9,10), and ischemic heart disease (10). The molecular mechanisms underlying changes in the activity of the L-type Ca 2ϩ channel remain under study for most pathologies.
The L-type Ca V 1.2 channel belongs to the molecular family of high voltage-activated Ca V channels. High voltage-activated Ca V 1.2 channels are hetero-oligomers composed of the main pore-forming Ca V ␣1 subunit non-covalently bound to the cytoplasmic Ca V ␤ auxiliary subunit, the EF-hand protein calmodulin (constitutively bound to the C terminus of Ca V ␣1), and the Ca V ␣2␦ subunit (11)(12)(13)(14)(15)(16). The full complement of auxiliary subunits is required to produce high voltage-activated Ca V 1.2 channels with the properties of the native channels. Ca V ␤ promotes the cell surface density of Ca V 1.2 channels through a high affinity interaction (17) in part by preventing its degradation by the ubiquitin/proteasome system (18). Co-expression of the Ca V ␣2␦ subunit with Ca V ␤-bound Ca V ␣1 increases peak current density and promotes channel activation at more negative voltages (19 -23). Although the molecular mechanism underlying this effect remains to be fully elucidated, the cell surface density of Ca V ␣2␦1 dictates the net Ca 2ϩ influx through L-type Ca 2ϩ channels (22). The Ca V ␣2␦1 subunit, encoded by CACNA2D1, is expressed in skeletal muscle (24) and in cardiac muscle (25) where it is the main isoform associated with Ca V 1.2 (25,26). The Ca V ␣2␦ proteins undergo complex co-and post-translational modifications. Endogenous Ca V ␣2 and Ca V ␦ are usually thought to be produced as a single protein that is proteolytically cleaved and then linked through strong disulfide bonds (27,28). Although Ca V ␣2␦ has been traditionally described as a type I transmembrane protein, it has been recently shown that Ca V ␣2␦ proteins associate with the plasma membrane through a glycosylphosphatidylinositol anchor attached to Ca V ␦ (29), although the functional relevance of this process remains to be fully established (30). Ca V ␣2␦ subunits are also heavily glycosylated. It is estimated that N-glycans contribute as much as Ϸ30 -50 kDa to the molecular mass of Ca V ␣2␦1 (26,27,31,32) making it a unique target for endoplasmic reticulum (ER) 3 glycoprotein quality control (33)(34)(35)(36).
Glycosylation is a form of co-and post-translational covalent modification that serves a variety of structural and functional roles in membrane and secreted proteins. The major classes of glycans being produced in eukaryotic cells are as follows: 1) Asn (N)-linked glycans attached to the nitrogen atom of asparagine side chains; 2) O-linked glycans attached to the hydroxyl oxygen of serine, threonine, tyrosine, hydroxylysine, or hydroxyproline side chains; and 3) C-linked glycans, a rare form of glycosylation, attached to a carbon on a tryptophan side chain. N-Glycosylation, one of the most abundant types of protein glycosylation (37), is initiated by the co-translational addition by the oligosaccharyltransferase of glucose 3 -mannose 9 -Nacetylglucosamine 2 core oligosaccharides to the Asn residue of the lumenally exposed consensus glycosylation site Asn-Xaa-(Ser/Thr) (NX(S/T)) where Xaa is any amino acid except proline (Pro), serine (Ser), and threonine (Thr) (38). The transfer of N-glycans to Asn-Xaa-(Ser/Thr) sites occurs on the lumenal side of the ER membrane while the protein moiety is being synthesized on ER-bound ribosomes; hence, only domains that are accessible to the ER lumen will receive N-glycans. Membrane glycoproteins remain anchored in the ER membrane with portion(s) either exposed to the ER lumen, embedded in the membrane, or within the cytoplasm. Subsequent trimming of glucose and mannose residues determines whether the polypeptide undergoes additional folding cycles or is targeted for the ER-associated degradation (ERAD) by retrotranslocation and ubiquitin proteasome-dependent proteolysis in the cytosol. This process is critical for protein biosynthesis, and abrogation of glycosylation causes embryonic lethality in mice (39).
Defective glycosylation of cardiac ion channels plays a role in multiple cardiac pathologies (40 -44). In this work, we have characterized the role of N-glycosylation on cell surface density and the function of the Ca V ␣2␦1 auxiliary subunit using mobility shift assays, cycloheximide pulse-chase analysis, confocal imaging, flow cytometry assays, and patch clamp recordings of recombinant Ca V 1.2 currents. Here, we showed that a single mutation at Asn-663 prevented the cell surface presentation and the function of Ca V ␣2␦1, although the protein remains strongly glycosylated. Mutations of other sites proved to alter channel function to a lesser extent. Simultaneous mutation of 6/16 consensus N-glycosylation sites in the extracellular portion of the Ca V ␣2␦1 protein curtailed protein stability and impaired channel function with a predominant role for Asn-348 and Asn-812. Furthermore, combining the N468Q mutation with N348Q and/or N812Q disturbed L-type channel function. Single mutations of the other Asn sites (out of the 16 tested) were without significant func-tional impact. Altogether, our data support a model where four Asn residues are essential to form functional L-type Ca V 1.2 currents.

Experimental Procedures
Recombinant DNA Techniques-The rabbit Ca V 1.2 (Gen-Bank TM accession number X15539) and the rat Ca V ␤3 (Gen-Bank TM accession number M88751) (45) were subcloned in commercial vectors under the control of the CMV promoter as described elsewhere (22,46). The primary sequence (1091 residues) of the rat brain Ca V ␣2␦1 clone (GenBank TM accession number NM_012919) (47) was subcloned in three vectors. Most experiments were performed with pmCherry-Ca V ␣2␦1-HA, where Ca V ␣2␦1 was subcloned in the pmCherry-N1 vector (Cederlane, Burlington, Ontario, Canada) between the SacI and SalI sites, and the hemagglutinin (HA) epitope (YPYDVPDYA) was inserted in the extracellular domain of Ca V ␣2 between Asp-676 and Arg-677 (22). Ca V ␣2␦1 was also subcloned in homemade vectors derived from the pCMV-Script vector and are referred to as pCMV-Ca V ␣2␦1 and pC2-Ca V ␣2␦1 in Fig. 2B. Except for Fig. 2B, the pmCherry-Ca V ␣2␦1-HA construct was used throughout.
Culture and Imaging of Mouse Cardiomyocytes-Experiments were approved by the Animal Protection Committee of the Montreal Heart Institute (protocol 2014-44-01) and were performed in accordance with the guidelines of the Canadian Council for Animal Care and the Guide for the Care and Use of Laboratory Animals 8th Edition (2011). Ventricular myocytes were isolated from neonate (22,48,49) or adult (50) CD-1 mice as described elsewhere. In the latter case, CD-1 male mice (5 months old) (Charles River Laboratories, St. Constant, Canada) were anesthetized with isoflurane. Hearts were quickly removed and placed on ice-cold Tyrode's solution containing 130 mM NaCl, 5.4 mM KCl, 1 mM MgCl, 0.33 mM Na 2 HPO 4 , 10 mM HEPES, 5.5 mM glucose, and 1 mM CaCl 2 , pH 7.4. Ventricles were isolated and homogenized at 4°C in a Tris-based solution containing a mix of protease inhibitors (Sigma), including 4-(2aminoethyl) benzenesulfonyl fluoride hydrochloride, aprotinin, bestatin, E-64, leupeptin, and 1 mM EDTA, pH 7.4 (50). Cells were fixed (2% paraformaldehyde, pH 7.4, 15 min, 4°C), blocked, and permeabilized (2% normal donkey serum, 0.1% Triton X-100, 60 min, at room temperature). Three washes with phosphate-buffered saline (PBS) followed each step. After overnight incubation at 4°C with primary anti-Ca V ␣2␦1 antibody (1:50) (Santa Cruz Biotechnology) in 1% donkey serum with 0.05% Triton X-100, cells were incubated with the secondary Alexa 488 antibody (Life Technologies, Inc.) (1:800) for 90 min at room temperature. Cells were stained with DAPI (1:1000) (Life Technologies, Inc.) for 10 min to identify the nucleus and with the wheat germ agglutinin-647 (WGA-647) (1:200) (Life Technologies, Inc.) to visualize cell membrane glycoproteins (51,52). WGA is a carbohydrate-binding protein of approximately 36 kDa that selectively recognizes sialic acid and N-acetylglucosaminyl sugar residues. Confocal fluorescent images were captured with a Zeiss LSM 710 confocal microscope system with a ϫ63/1.40 oil objective. The images were analyzed using FIJI software to delete background, subtract noise, and to produce co-localization pixel maps.
Live Imaging of HEKT Cells-HEKT cells stably transfected with Ca V ␤3 were transiently transfected simultaneously with pCMV-Ca V 1.2 and pmCherry-Ca V ␣2␦1-HA WT or NQ mutants. Cells were dissociated and seeded 6 h after transfection to obtain isolated cells for imaging. Exactly 24 h after transfection, cells were stained with the fluorescein isothiocyanate (FITC)conjugated mouse monoclonal anti-HA. Nuclei were stained with DAPI (1:1000) (Life Technologies, Inc.) in 1ϫ PBS for 45 min at 4°C. Confocal fluorescent images were captured with the same Zeiss LSM 710 confocal microscope used for cardiomyocyte imaging (see above).
Glycosidase Assays-Isolated mouse cardiomyocytes or transfected HEKT cells were solubilized in a radioimmunoprecipitation assay (RIPA) buffer (150 mM NaCl, 1.0% IGEPAL CA-630, 0.5% sodium deoxycholate, 0.1% SDS, and 50 mM Tris, pH 8.0) (Sigma) supplemented with a protease inhibitor mixture (Sigma). Cell lysates (20 g of proteins) were first incubated under denaturing conditions (0.5% SDS and 40 mM DTT) and then treated with 500 units of peptide-N-glycosidase F (PNGase F, New England Biolab) during 1 h at 37°C according to the manufacturer's instructions. Proteins were added to the Laemmli sample buffer in the presence of 0.4 mM 2-mercaptoethanol and electrophoresed on a 8% SDS-polyacrylamide gel alongside the Precision Plus Protein TM dual color standard (Bio-Rad). After electroblotting and blocking with 5% (w/v) skim milk for 30 min, the supported nitrocellulose membranes (Bio-Rad) were incubated with the anti-Ca V ␣2␦ (1:750, Alomone Labs, Jerusalem, Israel). Membranes were stripped and incubated with an anti-GAPDH as a loading control (1:10,000, Sigma) unless stated otherwise. Signal was detected with the Bio-Rad ECL chemiluminescent substrate. Blots were visualized with the ChemiDoc Touch documentation system (Bio-Rad). Molecular weights were estimated using Image Lab TM software by linear regression of standard molecular weight markers. The molecular mass of the Ca V ␣2␦1 protein in cardiomyocytes was calculated at 123 kDa. The calculated molecular mass of the mCherry-Ca V ␣2␦1-HA construct is 153 kDa. GAPDH migrated as a monomer close to 37 kDa in accordance with its calculated mass.
Cycloheximide Chase Assays-Stably transfected Ca V ␤3 cells were transiently transfected simultaneously with pCMV-Ca V 1.2 WT and pmCherry-Ca V ␣2␦1-HA WT or NQ mutants. Cells were incubated with 100 g/ml cycloheximide (Sigma) to block de novo protein synthesis 24 -36 h after transfection. At the indicated time points (0 h or no cycloheximide, 30 min and 1-4, 6, 10, and 24 h), cell lysates were fractionated on a 8% SDS-PAGE followed by immunoblotting to visualize Ca V ␣2␦1 (Alomone Labs, 1:750) and GAPDH (Sigma, 1:10,000). Protein density of Ca V ␣2␦1 in total lysates was estimated with Image-Lab 5.2 (Bio-Rad). It was expressed relative to GAPDH and normalized to the relative protein density of Ca V ␣2␦1-HA WT measured at time 0. The time course of degradation was measured in 3-5 experiments. Each symbol represents the mean Ϯ S.E. of the normalized protein density.
Isolation of the Plasma Membrane Fraction from Cardiomyocytes and HEKT Cells-Four different protein fractions (total cell lysates, cytosolic, total membrane, and plasma membrane fraction) were prepared according to a protocol published previously (50). Briefly, transfected HEKT cells cultured in 100-mm dishes were homogenized at 4°C in a Tris-based solution containing a mix of protease inhibitors (Sigma) and 1 mM EDTA, pH 7.4. The cell homogenate was aliquoted into three tubes. After a 2-h incubation period at 4°C with 1% (v/v) Triton X-100, the first tube was centrifuged at 10,000 ϫ g for 10 min to remove cell debris, nuclei, and mitochondria. The supernatant was kept as the total protein fraction (whole-cell lysates). The second tube was centrifuged at 200,000 ϫ g and 4°C for 20 min.
The supernatant is referred to as the cytosolic fraction. The pellet was resuspended in homogenizing buffer containing 1% (v/v) Triton X-100. After 30 min of incubation on ice, a second centrifugation was done at 200,000 ϫ g. The resulting supernatant is referred to as the total membrane protein fraction. The third tube was centrifuged at 10,000 ϫ g for 10 min. The supernatant obtained was centrifuged at 200,000 ϫ g and 4°C for 20 min. The pellet was resuspended in the homogenizing buffer containing 0.6 M KCl. Subsequent centrifugations were performed at 200,000 ϫ g and 4°C for 20 min to wash out KCl. The final pellet was resuspended in the homogenizing buffer and is considered to be enriched in plasma membrane proteins. Proteins were electrophoresed on an 8% SDS-polyacrylamide gel and blotted with the anti-Ca V ␣2␦ (Aviva System Biology 1:1000).
Flow Cytometry Assays-Flow cytometry experiments were conducted as described elsewhere (22). Stable Ca V ␤3 cells were transiently transfected simultaneously with pCMV-Ca V 1.2 WT and pmCherry-Ca V ␣2␦1-HA WT or mutants. To determine cell surface expression level of the mCherry-Ca V ␣2␦1-HA proteins, cells were harvested 24 h after transfection, washed in a 1ϫ PBS buffer, and stained with the FITC-conjugated mouse monoclonal anti-HA epitope tag antibody at 5 g/ml (Sigma) at 4°C for 30 min. To determine the total quantity of both intracellular and extracellular expression of the tagged proteins, cells were fixed and permeabilized using BD Cytofix/Cytoperm TM fixation/permeabilization solution kit (BD Biosciences) (22). Roughly 10,000 cells were counted using a FACSAria III SORP (Special Order Research Product) flow cytometer (BD Biosciences) at the flow cytometry facility hosted by the Department of Microbiologie, Infectiologie, and Immunologie at the Université de Montréal. The level of fluorescence detected with the IgG1-FITC isotype control murine (5 g/ml) or with the anti-HA FITC-conjugated antibody (5 g/ml) in HEKT untransfected cells was not significantly different from the fluorescence measured in the complete absence of fluorophore (22). Control conditions were carried out in triplicate with each series of experiments as follows: (a) untransfected Ca V ␤3 cells without anti-HA FITC-conjugated antibody; (b) untransfected Ca V ␤3 cells with the anti-HA FITC-conjugated antibody to assess the level of background staining; and (c) Ca V ␤3 cells transfected with pmCherry-Ca V ␣2␦1-HA WT. Expressing the mCherry-Ca V ␣2␦1-HA WT constructs in HEKT cells produced a significant 3-log increase in the FITC fluorescence (x axis) and mCherry fluorescence (y axis) on the two-dimensional plots (22).
Quantification of Steady-state Cell Surface Expression by Flow Cytometry Assays-Flow cytometry data were analyzed using the FlowJo software, version 10 (TreeStar, Ashland, OR) as described (22). Relative expression of Ca V ␣2␦1 was calculated based on ⌬mean fluorescence intensity (⌬MFI) for each fluorophore (mCherry or FITC) as explained elsewhere (22). Briefly, the positive cell gate (P2) and the negative cell gate (P3) were set manually. The fluorescence intensity within the region delineated by the P2 and P3 gates was displayed as cell count versus fluorescence intensity. The ⌬MFI for FITC was calculated by subtracting the FITC fluorescence density of the FITCnegative cells (P3) from the fluorescence density of the FITC-positive cells (P2). The same method was used to calculate the ⌬MFI for mCherry. Under our experimental conditions, the fluorescence intensity follows a normal distribution, hence the mean was equivalent to the median for all intents and purposes. ⌬MFI for FITC measured in intact non-permeabilized cells was used as a relative index of the steady-state cell surface density of the HA-tagged Ca V ␣2␦1. The ⌬MFI values for FITC were measured in permeabilized cells to confirm the accessibility of the HA epitope. It is also a valid estimation of the total protein density because the relative ⌬MFI values for FITC estimated in permeabilized cells are comparable with the relative ⌬MFI values for mCherry measured under the same conditions. ⌬MFI values were normalized to the maximum value measured the same day for mCherry-Ca V ␣2␦1-HA WT expressed under the same conditions to account for variations in the absolute fluorescence intensity of the anti-HA FITCconjugated antibody. The ⌬MFI values for FITC and mCherry obtained over the course of several months were pooled and are reported in Table 1, along with the number of triplicate experiments. The normalized ⌬MFI values for mCherry measured for each mutant in intact and permeabilized cells were not significantly different from one another (p Ͼ 0.05) suggesting that the cell permeabilization procedure did not distort significantly the relative fluorescence readout.
Patch Clamp Experiments in HEKT Cells-Whole-cell patch clamp experiments were carried out in isolated cells after transfection in HEKT Ca V ␤3 cells in the presence of the peGFP vector (0.2 g) as a control for transfection efficiency. Electrodes were filled with a solution containing (in mM) 140 CsCl, 0.6 NaGTP, 3 MgATP, 10 EGTA, 10 HEPES and titrated to pH 7.3 with NaOH. Cells were bathed in a modified Earle's saline solution (in mM) containing 135 NaCl, 20 TEACl, 2 CaCl 2 , 1 MgCl 2 , 10 HEPES and titrated to pH 7.3 with KOH. On-line data acquisition was achieved with the Axopatch 200-B amplifier (Molecular Devices, Sunnyvale, CA) connected with the PClamp software Clampex 10.5 through the Digidata 1440A acquisition system (Molecular Devices) (22). A series of 450-ms voltage pulses were applied from a holding potential of Ϫ100 mV at a frequency of 0.2 Hz, from Ϫ60 to ϩ70 mV at 5-mV intervals. Series resistance was compensated to ϳ85% after on line capacitive transient cancellation. Unless stated otherwise, whole-cell currents were sampled at 5 kHz and filtered at 1 kHz. PClamp software Clampfit10.5 was used for data analysis. Mid-potential of activation values (E 0.5,act ) were estimated from the peak I-V curves obtained for each channel composition and were reported as the mean of individual measurements Ϯ S.E (Table  2) (22,53). The free energy of activation was calculated using the mid-activation potential as shown in Equation 1, where z is the effective charge displacement during activation, and F is the Faraday constant (54). The r100 ratio is defined as the ratio of peak whole-cell currents remaining after a depolarizing pulse of 100 ms (I 100ms /I Peak ) and was used as an indicator of the inactivation kinetics. The pmCherry-Ca V ␣2␦1-HA construct was previously shown to carry the functional modulation of Ca V 1.2 currents (22). Each novel pmCherry-Ca V ␣2␦1-HA mutant was tested alongside the control WT construct (pCMV- to assess for internal consistency thus explaining the large sample size for the mCherry-Ca V ␣2␦1-HA WT construct. Experiments performed under the same conditions yielded peak current densities that could vary by as much as Ϯ35% between each series of experiments. This variation appeared to be essentially linked to minor changes in the cell density at the time of transfection. Data from all experiments performed under the same conditions over a period of 16 months were pooled, and biophysical properties are reported in Table 2. Experiments were performed at room temperature (18 -20°C).
Statistics-Results were expressed as mean Ϯ S.E. Tests of significance were carried out using the unpaired analysis of variance with the Tukey test embedded in the Origin 7.0 analysis software (OriginLab Corp., Northampton, MA). Data were considered statistically significant at p Ͻ 0.05.

Results
Cardiac Ca V ␣2␦1 Proteins Are Glycosylated-Ca V ␣2␦1 is largely believed to be the most heavily glycosylated protein within the cardiac L-type channel complex (32). Triple-color immunostaining of cultured mouse cardiomyocytes with wheat germ agglutinin (WGA), a plasma membrane marker that binds glycoproteins with high affinity, DAPI, a marker for the nucleus, and an anti-Ca V ␣2␦1 demonstrated that the glycoprotein Ca V ␣2␦1 co-localized with WGA (Fig. 1A). The expression of the endogenous Ca V ␣2␦1 in the sarcolemmal membraneenriched fraction in mouse cardiomyocytes was confirmed by membrane fractionation followed by SDS-PAGE. As seen, the major protein species migrated at an apparent molecular mass of 160 kDa in all fractions (Fig. 1B), which is similar to the mobility of the Ca V ␣2␦1 protein expressed in brain (29). Given that the calculated molecular mass of the rodent Ca V ␣2␦1 is 124 kDa, this suggests the native protein undergoes significant co-or post-translational modifications. To note, partially glycosylated species were not detected under our experimental conditions. Enzymatic deglycosylation carried out with PNGase F, an amidase that removes all saccharide moieties and reduced the electrophoretic mobility of the endogenous protein from 160 to 110 kDa (Fig. 1C). The 50-kDa decrease in the mobility of endogenous Ca V ␣2␦1 is compatible with the addition of N-glycans onto 10 -14 N-glycosylation sites (26,27,31).
Molecular Identification of N-Glycosylation Sites in Ca V ␣2␦1-By definition, N-linked glycans are attached to the nitrogen atom of an asparagine side chain within the Asn-Xaa-(Ser/Thr) consensus sequence, where Xaa is not a proline residue. It is estimated that at least two-thirds of those sites are likely to be N-glycosylated (55). Using this strict definition, we identified 16 putative glycosylation sites in the extracellular portion of the Ca V ␣2␦1 protein as follows: Asn-92, Asn-136, Asn-184, Asn-324, Asn-348, Asn-468, Asn-475, Asn-585, Asn-594, Asn-663, Asn-769, Asn-812, Asn-876, Asn-883, Asn-973, and Asn-986 in the rat isoform ( Fig. 2A and data not shown). Seven sites are conserved in the primary sequence of the human, rat, and mouse Ca V ␣2␦1 as follows: Asn-92, Asn-184, Asn-348, Asn-468, Asn-594, Asn-663, and Asn-812. The Asn-348 and Asn-468 sites are also conserved in Ca V ␣2␦2. Many Asn-Xaa-Ser sites are glycosylated inefficiently in vitro (56), whereas Asn-Xaa-Thr sites are usually efficiently glycosylated. Even though the presence of the Asn-Xaa-(Ser/Thr) site is necessary for the receipt of an N-glycan, transfer of the N-glycan to this site does not always occur, due to conformational or other constraints during glycoprotein folding (55). Enzymatic digestion with PNGase F carried out with whole-cell lysates from recombinant Ca V ␣2␦1 expressed in HEKT cells also demonstrated a reduction of 50 kDa in the electrophoretic mobility of Ca V ␣2␦1 (Fig. 2B), similar to the one observed above for the endogenous Ca V ␣2␦1 protein. The mobility shift assay also suggests that the Ca V ␦ protein may not be proteolytically FIGURE 1. A, endogenous Ca V ␣2␦1 proteins in mouse cardiomyocytes are glycoproteins. Endogenous Ca V ␣2␦1 in 24-h cultured mouse cardiomyocytes co-localized with wheat germ agglutinin 647 (WGA 647), a plasma membrane marker that displays a high affinity for glycoproteins. Ca V ␣2␦1 proteins were stained with the anti-Ca V ␣2␦1 as the primary antibody and Alexa 488-coupled secondary antibody. Scale bar corresponds to 10 m. The red channel was arbitrarily assigned to WGA, and the green channel was assigned to Ca V ␣2␦1. Nuclei are stained with DAPI (blue). Co-localization pixel maps of Ca V ␣2␦1 and WGA are shown in white and were produced using the co-localization finder plugin in FIJI. B, endogenous Ca V ␣2␦1 proteins in the sarcolemmal membrane fraction of mouse cardiomyocytes migrate at 160 kDa. Four different protein fractions (total (WCL), cytosolic (Cyto), total membrane (Tot mb), and sarcolemmal membrane (Sarc mb)) were isolated from ventricles of adult CD-1 mice (50). Proteins were electrophoresed on an 8% SDS-polyacrylamide denaturating gel, transferred to a nitrocellulose membrane, and probed with an anti-Ca V ␣2␦1 (Aviva System Biology). The membrane was probed, after stripping, with antipan-cadherin (Invitrogen 1:5000) as a quality control for the fractionation process. It is worth noting that the 160-kDa protein is the dominant species in whole-cell lysates. Each lane was loaded with 10 g of proteins. C, PNGase F-mediated deglycosylation of the N-linked sugars in endogenous Ca V ␣2␦1 proteins from mouse cardiomyocytes. Total whole-cell proteins isolated from ventricles of adult CD-1 mice were denatured 20 min at 60°C before incubation in the absence (Ϫ) or presence (ϩ) of PNGase F during 1 h at 37°C. Each lane was loaded with 20 g of proteins. Proteins were electrophoresed on an 8% SDS-polyacrylamide denaturing gel, transferred to a nitrocellulose membrane, and probed with an anti-Ca V ␣2␦1 (Alomone Labs) (top panel) and anti-GAPDH (bottom panel) as a loading control.
cleaved under our experimental conditions (57,58). The molecular determinants responsible for N-type glycosylation were investigated using a mutational analysis. Multiple Asn to Gln mutants were produced in the pmCherry-Ca V ␣2␦1-HA construct that was previously shown to be expressed at the plasma membrane and fully functional (22). The glycosylation status of multiple constructs 6xNQ (N92Q/N184Q/N348Q/N594Q/N812Q/ N876Q) that includes the six Asn sites that were predicted to be the most likely to be glycosylated and the 14xNQ construct (N92Q/N136Q/N184Q/N348Q/N468Q/N585Q/N594Q/ N663Q/N769Q/N812Q/N876Q/N883Q/N986Q/N1066Q) that includes the 14 Asn sites with a likelihood above 0.5 (59) were first investigated. Western blots produced with whole-cell lysates showed that pmCherry-Ca V ␣2␦1-HA WT migrated as a doublet with a faint band at Ϸ200 kDa and a stronger band Ϸ175 kDa (Fig. 2C). The 6xNQ and the 14xNQ proteins produced a band pattern of significantly weaker intensity with protein mobility reduced by Ϸ25 and Ϸ45 kDa, respectively (Fig. 2C), suggesting that acquisition of N-glycans was not completely impaired in the 6xNQ and the 14xNQ mutants. Indeed, enzymatic digestion with PNGase F further reduced the electrophoretic mobility and produced similar migration profiles for the mCherry-Ca V ␣2␦1-HA WT, 6xNQ, and the 14xNQ proteins. Altogether, this observation supports the view that some of the Asn residues mutated in the 6xNQ construct (Asn-92, Asn-184, Asn-348, Asn-594, Asn-812, and/or Asn-876), some of the additional residues substituted in the 14xNQ construct (Asn-136, Asn-468, Asn-585, Asn-663, Asn-769, Asn-883, Asn-986, and/or Asn-1066), and possibly the two remaining residues (Asn-475 and Asn-973) are acquiring N-glycans either co-translationally or post-translationally.
Disrupting Six Asn Sites Impaired Steady-state Surface Density of Ca V ␣2␦1-To evaluate whether the partially glycosylated forms reach the plasma membrane, we quantified the cell surface fluorescence of the mCherry-Ca V ␣2␦1-HA WT and NQ constructs in two-color flow cytometry assays. In this construct, the mCherry fluorescence is constitutive. The fluorescence of the FITC-conjugated HA antibody was shown to be proportional to the fraction of proteins present at the cell surface because the HA epitope is located in the extracellular portion of Ca V ␣2␦1 (22). The fluorescence intensity ⌬MFI for FITC measured in intact cells, observed as a rightward shift in the fluorescence intensity on the x axis of the two-dimensional plots, provides a reliable index of the steady-state cell surface density of Ca V ␣2␦1. The mCherry epitope expressed at the C terminus of the construct, observed as an increase in the fluorescence intensity seen on the y axis of the two-dimensional plots, served as a marker for total protein expression. The twocolor assay thus provided a quick and reliable readout of protein expression. Control experiments carried out with the mCherry-Ca V ␣2␦1 WT control construct that was not HA-tagged confirmed the specificity of the FITC antibody in these series of experiments (Fig. 3A). The fluorescence histograms for the corresponding experiment were reported to the right of the contour plots, whereas the averaged mean fluorescence intensity values (⌬MFI) obtained from Ն3 distinct experiments are shown in Fig. 3C. As seen, the fluorescence intensity ⌬MFI for FITC in intact nonpermeabilized cells was strong for the WT and the 4xNQ construct but sharply decreased from the 5xNQ to the 13xNQ constructs (data not shown). These constructs produced proteins that were almost absent from the cell surface. Nonetheless, a slightly different version of the 6xNQ mutant (N92Q/N184Q/N468Q/N594Q/N876Q/N986Q) was present at the cell surface at a density similar to the 4xNQ mutant (data not shown). Assays were also conducted after cell permeabilization, and the fluorescence histograms shown alongside confirmed the accessibility of the HA epitope. Altogether, these results suggest the following: 1) mutations of consensus N-glycosylation sites are associated with decreased steady-state cell surface density of Ca V ␣2␦1; 2) mutations of consensus N-glycosylation sites impaired total protein density; and 3) N-glycosylation sites may not all be functionally equivalent.
Disrupting Asn Sites Impairs Channel Function-Mutating the consensus N-linked glycosylation reduced the steady-state cell surface density of the Ca V ␣2␦1 protein. Its impact on the L-type Ca V 1.2 channel function was explored after recombinant expression of Ca V 1.2 with mCherry-Ca V ␣2␦1-HA multi-ple NQ mutants in stable Ca V ␤3 cells. As reported before (22), co-expression of pmCherry-Ca V ␣2␦1-HA WT with Ca V 1.2 WT in stable Ca V ␤3 cells stimulated whole-cell peak current densities from Ϫ3 Ϯ 1 pA/pF (n ϭ 35) (no insert in the pmCherry vector) to Ϫ30 Ϯ 1 pA/pF (n ϭ 231) in the presence of Ca V ␣2␦1 WT (Fig. 3, B and D). The increase in peak current densities was associated with an Ϸ Ϫ15-mV leftward shift in the activation potential of Ca V 1.2 from E 0.5, act ϭ 8 Ϯ 2 mV (n ϭ 35) (no Ca V ␣2␦1) to E 0.5, act ϭ Ϫ9.6 Ϯ 0.1 mV (n ϭ 231) (with mCherry-Ca V ␣2␦1-HA). As seen, co-expression with mCherry-Ca V ␣2␦1-HA mutants containing 4 (4xNQ), 6 (6xNQ), and 13 (13xNQ) mutations yielded voltage-activated Ca 2ϩ currents. Mutating four sites did not appreciably alter whole-cell peak current density, but mutations of two additional consensus sites in the 6xNQ mutant (N92Q/ N184Q/N348Q/N594Q/N812Q/N876Q) was sufficient to significantly decrease by 6-fold the peak current density of Ca V 1.2 currents. Furthermore, peak current densities measured with the 13xNQ mutant were not statistically different from Ca 2ϩ currents obtained in the absence of Ca V ␣2␦1 (data not shown). These results suggest that residues Asn-184 and Asn-812 could be among the most critical residues in carrying channel modulation. Only a Few Asn Residues Contribute to Ca V ␣2␦1 Function-Cell surface density, protein stability, and channel function were significantly impaired in the 6xNQ construct. To identify the individual contribution of each residue to the functional response, reverse Gln to Asn mutations were introduced in the 6xNQ construct. As seen, reinstating the Asn-812 site, with the "6xNQ ϩ Q812N" mutant, nearly restored the steady-state cell surface density of Ca V ␣2␦1 and L-type channel function (Fig. 4,  A and B). Reintroducing Asn-348 with the "6xNQ ϩ Q348N" mutant was also seen to significantly improve cell surface density and channel function. These data suggest that N348Q could account for the large decrease in the cell surface density of the 4xNQ mutant. In both cases, the increase in the relative cell surface density of Ca V ␣2␦1 (Fig. 4C) was correlated with an augmentation of the peak current density as compared with the 6xNQ mutant (Fig. 4D). In addition, the mean ⌬MFI for FITC in permeabilized cells (an index of total cell density) increased in these two reverse mutants suggesting that protein stability and/or synthesis was also improved as compared with the 6xNQ mutant (Fig. 4C). The individual impact of each Asn site was finally investigated in single point mutations. Sixteen single Asn to Gln mutations were tested (N92Q; N136Q; N184Q; N324Q; N348Q; N468Q; N475Q; N585Q; N594Q; N663Q; N769Q; N812Q; N812A; N876Q; N883Q; N973Q; and N986Q) ( Fig. 5 and Table 1). Most single mutations only caused small changes in the relative fluorescence intensity at the cell surface without significant change in channel gating, suggesting that these single mutants reached the cell surface in their native conformation. One can suppose that small changes in the surface fluorescence could result from minor alterations in the protein trafficking or protein conformation. Nine mutations (N136Q; N324Q; N475Q; N585Q; N594Q; N769Q; N876Q; N883Q; and N986Q) produced fluorescent patterns for FITC and mCherry not significantly different (p Ͼ 0.05) than the wild-type construct suggesting that neither surface density nor protein stability was affected by these single mutations. Three single mutations mCherry-Ca V ␣2␦1-HA N92Q, mCherry-Ca V ␣2␦1-HA N184Q, and mCherry-Ca V ␣2␦1-HA N973Q produced proteins that yielded slightly smaller fluorescent signals (by Ϸ15-20%) than the control mCherry-Ca V ␣2␦1-HA WT construct (p Ͻ 0.05). However, all these above-mentioned single NQ mutants stimulated peak currents to the same extent as the wild-type construct (p Ͼ 0.05) ( Table 2).
Four single point mutations (N348Q; N468Q; N663Q; and N812Q) significant decreased the fluorescence at the cell surface (p Ͻ 0.001). The strongest impact produced by a single mutation was obtained with N663Q that eradicated cell surface fluorescence and channel function ( Fig. 5 and Tables 1 and 2). N812Q and N812A also produced proteins that significantly decreased cell surface fluorescence. Data from the flow cytometry assays were corroborated with confocal images captured with live cells stained with the FITC-conjugated anti-HA tag antibody. Cell surface fluorescence intensity decreased sharply for the 6xNQ mutant and was noticeably weaker for the single N812Q mutant than for the wild-type construct (Fig. 6). In agreement with the fluorescence data, N812Q generated currents that were twice larger than the 6xNQ mutant but Ϸ3 times smaller than currents produced with the wild-type construct (p Ͻ 0.01) ( Table 2).
The potentially additive effect of these residues (save for N663Q that was already non-functional on its own) was investigated in double mutants. Pairing Asn sites in different combinations, to form double mutants N348Q/N468Q, N348Q/ N812Q, and N468Q/N812Q, produced proteins that significantly reduced the steady-state cell surface density of Ca V ␣2␦1 and modulation of Ca V 1.2 currents (Fig. 7, Tables 1 and 2; and data not shown). Pairing N92Q with either N348Q, N468Q, or N812Q as one of the partners yielded voltage-activated currents that were roughly 50% lower than produced with the wild-type construct. In contrast, other mCherry-Ca V ␣2␦1-HA double NQ mutants (N92/N184Q, N92Q/ N594Q, N136Q/N184Q, N136Q/N769Q, and N594Q/N876Q) produced whole-cell currents similar to the mCherry-Ca V ␣2␦1-HA WT construct (p Ͼ 0.05). The series of wild-typelike double mutants include N136Q/N184Q previously shown to prevent the subunit-mediated regulation of Ca V 2.2 currents (60). These results suggest that Asn-348, Asn-468, Asn-663, and Asn-812 in Ca V ␣2␦1 play unique roles in the modulation of Ca V 1.2.
Mutations of these sites produced Ca V ␣2␦1 proteins with impaired N-glycosylation. Mobility shift assays were carried out before and after digestion with PNGase F in three separate series of experiments as follows: with double mutants N136Q/ N184Q, N348Q/N468Q, N348Q/N812Q, and N468Q/N812Q (Fig. 8A), multiple mutants that were not detected at the membrane N348Q/N468Q/N812Q and N92Q/N184Q/N348Q/ N468Q/N594Q/N812Q (Fig. 8B), and single N663Q (Fig. 8C). Under control conditions, there was a small but significant decrease in the protein mobility (Ͻ10 kDa) for the double mutants when compared with the WT construct and a smaller one with single mutants, although this was not always clearly evident. Digestion with PNGase F decreased the electrophoretic mobility of N663Q, as well as the double, triple, and sextuple mutants, respectively by 48, 45, 40, and 30 kDa, respec-

TABLE 1 Relative fluorescence intensity ⌬MFI for Cherry-Ca v ␣2␦1-HA WT and mutants
Ca v 1.2 WT was co-expressed in stable Ca v ␤3 HEKT cells with pmCherry-Ca v ␣2␦1-HA WT or mutant using a 1:1 DNA ratio. Flow cytometry experiments were conducted to determine cell surface expression levels of tagged proteins, and fluorescence intensity was measured with the FlowJo software as described under "Experimental Procedures." Relative expression of Ca v ␣2␦1 was calculated based on ⌬MFI estimated for each fluorophore (mCherry or FITC). ⌬MFI for FITC measured in intact non-permeabilized cells was used as an index of the cell surface density of the HA-tagged Ca v ␣2␦1, and ⌬MFI values for FITC measured in permeabilized cells reflect the total protein expression (cell surface and intracellular protein density). The ⌬MFI values for the mCherry-Ca v ␣2␦1-HA mutants were pooled and normalized to the maximum value obtained for pmCherry-Ca v ␣2␦1-HA WT that was expressed under the same conditions and measured the same day. The total number of experiments is provided in parentheses. The ⌬MFI values for FITC measured in permeabilized cells were mostly similar to the ⌬MFI for mCherry measured in intact and permeabilized cells. Furthermore, the ⌬MFI values for mCherry measured in intact and permeabilized cells were found to be within experimental error suggesting that cell permeabilization did not significantly alter the protein structure. Statistical analysis was carried out against the ⌬MFI for FITC measured with pmCherry-Ca v ␣2␦1-HA WT (* p Ͻ 0.05; ** p Ͻ 0.01). FEBRUARY 26, 2016 • VOLUME 291 • NUMBER 9

.2/Ca v ␤3 with Ca v ␣2␦1 WT and mutants
Ca v 1.2 WT was co-expressed in stable Ca v ␤3 cells with pmCherry-no insert or pmCherry-Ca v ␣2␦1-HA WT or mutant using a 1:1 DNA ratio. Biophysical parameters were measured in the presence of 2 mM Ca 2ϩ as described elsewhere (17,22). Activation properties (E 0.5,Խact and ⌬G act ) were estimated from the mean I-V relationships and fitted to a Boltzmann equation. The data are shown with the mean Ϯ S.E. of the individual experiments and the number of experiments appears in parentheses. N.D. not determined because of a poor signal to noise ratio. Statistical analysis was carried out against the values obtained in the presence of mCherry-Ca v ␣2␦1-HA WT (* p Ͻ 0.05; ** p Ͻ 0.01).
tively. These results confirm that the simultaneous mutation of these Asn sites significantly affected the glycosylation status of Ca V ␣2␦1, although the Ca V ␣2␦1 protein remains strongly N-linked glycosylated. Together, these data show that the Ca V ␣2␦1 protein is heavily glycosylated on many if not all of the 16 Asn sites because the simultaneous mutation   FEBRUARY 26, 2016 • VOLUME 291 • NUMBER 9 JOURNAL OF BIOLOGICAL CHEMISTRY 4837 of the 16 sites eliminated the formation of the glycosylated protein (Fig. 8D).

N-Glycosylation of the Cardiac L-type Channel Complex
Protein Stability/Synthesis Was Impaired in Asn Mutants-The decrease in the cell surface density of the double mutants N348Q/N812Q and N468Q/N812Q was accompanied by a 50 -60% decrease in the FITC fluorescence under permeabilized conditions suggesting that protein stability was severely altered. Chase assays were carried out in the presence of cycloheximide (a blocker of de novo protein synthesis) to document the time course of protein degradation. Cycloheximide was added 24 h after transfection, and total cell lysates were collected at different time points. Protein density was estimated from Western blots relative to the loading control GAPDH and normalized using the protein density of the WT construct at time 0 (Fig. 9, A-C). In all cases, the major band formed by the mCherry-Ca V ␣2␦1-HA WT construct disappeared with a halflife estimated at 2.8 Ϯ 0.5 h (n ϭ 7) (Fig. 9D). As compared with the WT construct, the relative protein density of the double mutants N348Q/N812Q and N468Q/N812Q were significantly lower even at time 0. The degradation kinetics were also slightly faster with t1 ⁄ 2 Ϸ1.2 Ϯ 0.8 h (n ϭ 2) and t1 ⁄ 2 ϭ 1.8 Ϯ 0.5 h (n ϭ 3), respectively. These data suggest that N-glycosylation at these sites is required for protein translation or else that kinetics of degradation of NQ mutants are faster than the kinetics of pro-tein synthesis, an observation that was also reported for glycosylation-defective mutations of the type 1 transmembrane auxiliary subunit KCNE1 (43).

Discussion
16 Asn Sites Account for N-type Glycosylation of Ca V ␣2␦1-N-Linked glycosylation is one of the most common post-translational modifications known to influence the turnover and the stability of cardiac ion channels (61)(62)(63)(64)(65). With the cardiac Ca V 1.2 macromolecular complex, Ca V ␣2␦1 is the most heavily glycosylated protein (13) with N-glycans increasing the apparent molecular mass by about 50 kDa. The fully glycosylated form was found to be the dominant protein species of the endogenous Ca V ␣2␦1 found in the plasma and associated caveolae membranes of isolated cardiomyocytes. In this work, we have addressed the role of N-glycosylation in the protein density, steady-state cell surface levels, and the function of the Ca V ␣2␦1 auxiliary subunit using mobility shift assays, cycloheximide pulse-chase analysis, flow cytometry assays, and patch clamp recordings of recombinant Ca V 1.2 currents. The 16 consensus N-type glycosylation sites were characterized after single or multiple mutations of Asn to Gln. By combining fluorescence and functional assays, we demonstrated that multiple glycosylation-defective mutants reduced the steady-state cell surface density, decreased total protein density, and diminished protein stability of Ca V ␣2␦1. The drop in the cell surface expression of Ca V ␣2␦1 in turn significantly impaired peak current density and activation gating of the L-type Ca V 1.2 channel.
Enzymatic digestion with PNGase F of the native and recombinant Ca V ␣2␦1 reduced by 50 kDa the electrophoretic mobility of the protein. A similar mobility shift was observed when comparing the migration profile of the WT and the 16xNQ construct demonstrating that many if not all the 16 Asn sites are required to account for the N-glycosylated state of the Ca V ␣2␦1 protein. Interestingly, the recently published three-dimensional structure of the skeletal muscle Ca V 1.1 channel complex is compatible with the proposition that most 16 N-glycan sites are glycosylated (66). The nature of the carbohydrate chain modifications that the protein undergoes from the endoplasmic reticulum to the post-Golgi compartments was not investigated in this work. We used throughout PNGase F, an amidase that cleaves between the innermost N-acetylglucosamine and Asn residues of high mannose, hybrid, and complex oligosaccharides from N-linked glycoproteins, thus stripping all glycans from the protein. Protein biogenesis further includes trimming of glucose and mannose residues by glycosidases and addition of new residues via glycosyltransferases in the ER and, to a great extent, in the Golgi. In the Golgi, high mannose N-glycans can be converted to a variety of complex and hybrid forms that are unique to each protein (67). The Ca V ␣2␦1 protein thus contains multiple glycosylation sites that may be modified with any of the three classes of N-linked glycans. Adding to the complexity, different units of the same Ca V ␣2␦1 glycoprotein may have different glycan structures attached to the identical Asn site. From the mobility shift assays conducted with the multiple mutants, it can be argued that many of these 16 sequons are modified by N-glycans even NXS sequons with negatively charged or hydrophobic amino acids at the X-position (such as Asn-136 and Asn-184) that are considered to be poorer substrates for the oligosaccharyltransferase complex (43,56). The N-glycan modification of many sequons appears to individually contribute 3-5 kDa to Ca V ␣2␦1. Although this is often difficult to detect visually for single mutations, it can be seen more clearly with double mutants.
Mutation of Asn-663 Prevents Cell Surface Density of Ca V ␣2␦1-The single mutation of Asn-663 (N663Q) prevented the detection of the protein at the cell surface by immunofluorescence and channel modulation. Asn-663 is located close to the 9-residue HA epitope that was inserted after Asp-676 in the primary structure of Ca V ␣2␦1. However, substitution of the asparagine for the glutamine residue did not prevent protein synthesis as the protein was expressed with the expected molecular mass. Furthermore, the mCherry and the HA epitope of this construction were fluorescently labeled and detected in intact and permeabilized cells, respectively. Every other single Asn mutation within the 16 sites did not significantly affect cell surface density or protein function thus making Asn-663 one of the most critical single sites for protein expression. Asn sites in the 6xNQ mutant prevented protein detection at the surface and impaired modulation of Ca V 1.2 currents. Double mutations pairing N348Q, N812Q, and/or N468Q were sufficient to prevent the detection of Ca V ␣2␦1 at the cell surface and abolished the subunit-mediated stimulation of Ca V 1.2 currents. Other double Asn mutations, including the N136Q/N184Q mutant (60), did not affect cell surface density or protein function. One reason for this discrepancy may lie FIGURE 9. Multiple Asn mutations may impair protein stability. HEKT cells were transiently transfected with pmCherry-Ca V ␣2␦1-HA WT and the glycosylation mutants N348Q/N812Q (A), N468Q/N812Q (B), and N812Q (C). Exactly 24 h after transfection, subconfluent cells were incubated with 100 g/ml cycloheximide. At the indicated time points, cell lysates were prepared and fractionated by SDS-PAGE (8%). Ca V ␣2␦1 and GAPDH proteins were respectively probed with anti-Ca V ␣2␦1 (Alomone Labs) and anti-GAPDH. Please note that we loaded 2ϫ more proteins in the wells for the double mutants to visualize their time course alongside the WT construct. In particular, each lane for Ca V ␣2␦1-HA WT was loaded with 5 g of proteins in A and B, and 10 g of proteins were loaded for N348Q/N812Q and N468Q/N812Q. C, 10 g of proteins were loaded for Ca V ␣2␦1-HA WT and N812Q. D, protein density was estimated relative to the density of the GAPDH band and normalized to the protein density for the wild-type construct at time 0. Protein band intensities were quantified by densitometry using ImageLab (Bio-Rad) software at a single exposure selected for clear bands without saturation. Averaged data points were fitted with a monoexponential decay function. The half-life was t1 ⁄2 ϭ 2.8 Ϯ 0.5 h (n ϭ 7) for pmCherry-Ca V ␣2␦1-HA WT; t1 ⁄2 ϭ 1.2 Ϯ 0.8 h (n ϭ 2) for N348Q/N812Q; t1 ⁄2 ϭ 1.8 Ϯ 0.5 h (n ϭ 3) for N468Q/N812Q; and t1 ⁄2 ϭ 1.9 Ϯ 0.7 h (n ϭ 3) for N812Q. As seen, the data points for the decay of N812Q and N468Q/N812Q are superimposed with similar relative densities, whereas protein density for N348Q/N812Q was lower at time 0. in the differences in the mode of regulation of Ca V 1.2 and Ca V 2.2 channels by Ca V ␣2␦1. To limit the number of double mutants to be tested (120 possible double mutants for 16 sequons), our strategy was to produce and characterize multiple mutants that impacted on channel function. Multiple mutants produced with either N348Q, N468Q, or N812Q showed a significant decrease in cell surface density and protein function, whereas every single mutation pairing N663Q was not functional. It is impossible to know at this time whether N-glycans at these sites interact with each other or with the poreforming subunit. Current algorithms and the recent three-dimensional structure of the skeletal Ca V 1.1 channel complex (66) are identifying important structural domains but are not predicting the spatial orientation of the 16 N-glycans with each other and other proteins in the channel.
Protein Stability Was Decreased in Glycosylation-defective Mutants of Ca V ␣2␦1-The molecular and cellular pathways responsible for the compromised steady-state cell surface expression of the glycosylation-defective Ca V ␣2␦1 proteins remain to be fully elucidated. The decreased surface levels could result from impaired translational rate (68,69), ER folding yield, trans-Golgi cargo sorting, increased degradation, and/or a combination of these processes. The relative contribution of lectin chaperones, such as calnexin and calreticulin, to protein biogenesis and the nature of the key enzymes responsible for degradation within the ubiquitin-proteasome system of ERAD (68, 70 -72) also remain to be identified. Outstanding issues include the identification of the quality control networks that are affected by these specific glycosylation sites (73,74) and the potential cross-talk between ERAD and autophagy, the two major cellular degradative pathways (75). Finally, there is always the question of overexpressing cardiac proteins in a model cell to study translation events. These are important questions that currently go beyond the scope of this work. Protein assays herein reported nonetheless support the view that protein stability and/or protein synthesis was impaired in Asn mutants suggesting that N-glycosylation is a co-translational event (43). The observation that partially glycosylated forms of Ca V ␣2␦1 were notably absent from whole-cell homogenates prepared with recombinant cells as well as isolated cardiomyocytes further argues for this scenario.
L-type Channel Modulation by Ca V ␣2␦1-Up-regulation of L-type currents requires robust cell surface density of Ca V ␣2␦1 in cardiomyocytes (19) and in HEKT cells (22). Alterations in the cell surface density of Ca V ␣2␦1 caused by mutating glycosylation sites (our study), arrhythmogenic mutations (22), mutations within the "von Willebrand factor" structural domain (76,77), or following pharmacological modulation (e.g. gabapentin) (76,78,79) were shown to decrease channel function by altering the surface levels of Ca V ␣2␦1 (76,77). With the exception of the so-called R-domain (80), these manipulations altered the function of Ca V ␣2␦1 (and by extension the function of voltage-gated currents) through a decrease in the membrane expression of Ca V ␣2␦1. By analogy with other type 1 transmembrane regulatory subunits of voltage-gated ion channels, Ca V ␣2␦1 could interact with the pore-forming subunit either within the membrane through the voltage sensing domain (as KCNE with Kv7/KCNQ1 channel (81)) and/or from the exter-nal portion of the protein by interacting with the external pore domain (77). The optimal ratio for channel modulation remains to be established. By comparison, a single high affinity intracellular binding site for Ca V ␤ onto the I-II linker of the Ca V ␣1 subunit from high voltage-activated Ca V 1 and Ca V 2 channels has been identified (17,46,(82)(83)(84). Whether a single Ca V ␣2␦1 subunit could interact with two Ca V ␣1 or whether the L-type Ca V 1.2 channel complex (85) can accommodate two or more Ca V ␣2␦1 subunits remains a question for debate. Nonetheless, mutations affecting the expression of Ca V ␣2␦1 is likely to influence Ca 2ϩ balance in cardiomyocytes (86 -88) as in arterial smooth muscle cells (4) by virtue of controlling the activity of L-type Ca V 1.2 channels (22).
Recent technical advances in glycoprotein crystallography suggest that the more mobile N-glycans on cell surface receptors could guide the partner ligand to its binding site and prevent irregular protein aggregation by covering oligomerization sites away from the ligand-binding site (91). Beyond their role in protein biogenesis and/or stability (37), N-linked glycans promote interactions with cell adhesion proteins and signaling molecules (40,89) as it is widely understood for members of the integrin family (90). Glycans could also contribute to channel modulation by altering the surface potential sensed by the gating machinery (92) and/or by modifying conformational changes regulating cooperative subunit interactions during channel activation (93). The Asn-812 site is a choice candidate for such a mechanism. The N812Q mutant was expressed at the cell surface to the same extent as 4xNQ mutant, yet it caused a more extensive decrease in channel function. It can be speculated that N-glycans at the Asn-812 site contribute to the functional interaction with the pore-forming subunit of the Ca V 1.2 channel, either through direct protein-protein interaction and/or by promoting a favorable gating conformation (31). Most structural models predict that the extracellular domain is quite disordered, making it impossible to predict the relative orientation and/or interaction of each N-glycan chain within the Ca V 1.2 channel complex. The nature of the protein-protein interaction (either direct or through a secondary partner), the sites responsible for this interaction, and well as the affinity of the interaction will await further structural characterization of the cardiac Ca V 1.2 channels. At this time, three-dimensional structures of the purified cardiac Ca V 1.2 channel complex position the extracellular portion of Ca V ␣2␦1 on top of the channel complex at a resolution that prevents identifying interaction domains (32), although the recent three-dimensional structure of the skeletal muscle Ca V 1.1 channel complex suggests that the VWA domain in Ca V ␣2␦1 may be directly interacting with the voltage-sensing region of the pore-forming Ca V ␣1 subunit (66). Nonetheless, it is becoming quite evident that biological networks exploit cell-surface glycans to coordinate membrane protein complexes (94). Defects in the glycosylation of type I transmembrane auxiliary subunit Ca V ␣2␦1 could hence trigger ventricular and atrial arrhythmias (22,95) by decreasing the fraction of functional L-type Ca V 1.2 channels. Hence, elucidating the cellular processes controlled by glycosylation contributes to furthering our understanding of most biological systems and, in particular, voltage-gated cardiac ion channels.
Author Contributions-M. P. T. produced the single and multiple Asn mutants, performed and analyzed the vast majority of the flow cytometry experiments, carried out the cycloheximide chase assays, and stained the HEKT cells for confocal microscopy. B. B. isolated the mouse cardiomyocytes, conducted patch clamp experiments, isolated the sarcolemmal membrane fractions from mouse cardiomyocytes and the plasma membrane fraction from recombinant HEKT cells, as well as carried out the Western blots in Fig. 1B and Fig. 2A. J. B. produced some constructs, and performed the experiments shown in Fig. 2C. E. S. conducted flow cytometry and patch clamp experiments for a few mutants. S. L. supervised the analysis of the flow cytometry experiments. C. F. supervised the isolation and the culture of the mouse cardiomyocytes. L. P. designed and coordinated the study, interpreted the data, and wrote the manuscript. All authors reviewed the results and approved the final version of this manuscript.