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Ca²⁺ Current and Charge Movements in Skeletal Myotubes Promoted by the ß-Subunit of the Dihydropyridine Receptor in the Absence of Ryanodine Receptor Type 1

Chris A. Ahern^*, David. C. Sheridan^*, Weijun Cheng^*, Lindsay Mortenson^*, Priya Nataraj^*, Paul Allen, Michel De Waard and Roberto Coronado^*

^* Department of Physiology, University of Wisconsin School of Medicine, Madison, Wisconsin 53706; Department of Anesthesiology, Brigham and Women's Hospital, Boston Massachusetts 02115; and CEA, INSERM EMI 9931, UJF, CNRS, Laboratoire de Canaux Ioniques et Signalisation, 17 rue des Martyrs 38054 Grenoble, France

Correspondence: Address reprint requests to Roberto Coronado, Dept. of Physiology, University of Wisconsin, 1300 University Avenue, Madison, WI 53706. Tel.: 608-263-7487; Fax: 608-265-5512; E-mail: coronado{at}physiology.wisc.edu.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
The ß-subunit of the dihydropyridine receptor (DHPR) enhances the Ca²⁺ channel and voltage-sensing functions of the DHPR. In skeletal myotubes, there is additional modulation of DHPR functions imposed by the presence of ryanodine receptor type-1 (RyR1). Here, we examined the participation of the ß-subunit in the expression of L-type Ca²⁺ current and charge movements in RyR1 knock-out (KO), ß1 KO, and double ß1/RyR1 KO myotubes generated by mating heterozygous ß1 KO and RyR1 KO mice. Primary myotube cultures of each genotype were transfected with various ß-isoforms and then whole-cell voltage-clamped for measurements of Ca²⁺ and gating currents. Overexpression of the endogenous skeletal ß1a isoform resulted in a low-density Ca²⁺ current either in RyR1 KO (36 ± 9 pS/pF) or in ß1/RyR1 KO (34 ± 7 pS/pF) myotubes. However, the heterologous ß2a variant with a double cysteine motif in the N-terminus (C3, C4), recovered a Ca²⁺ current that was entirely wild-type in density in RyR1 KO (195 ± 16 pS/pF) and was significantly enhanced in double ß1/RyR1 KO (115 ± 18 pS/pF) myotubes. Other variants tested from the four ß gene families (ß1a, ß1b, ß1c, ß3, and ß4) were unable to enhance Ca²⁺ current expression in RyR1 KO myotubes. In contrast, intramembrane charge movements in ß2a-expressing ß1a/RyR1 KO myotubes were significantly lower than in ß1a-expressing ß1a/RyR1 KO myotubes, and the same tendency was observed in the RyR1 KO myotube. Thus, ß2a had a preferential ability to recover Ca²⁺ current, whereas ß1a had a preferential ability to rescue charge movements. Elimination of the double cysteine motif (ß2a C3,4S) eliminated the RyR1-independent Ca²⁺ current expression. Furthermore, Ca²⁺ current enhancement was observed with a ß2a variant lacking the double cysteine motif and fused to the surface membrane glycoprotein CD8. Thus, tethering the ß2a variant to the myotube surface activated the DHPR Ca²⁺ current and bypassed the requirement for RyR1. The data suggest that the Ca²⁺ current expressed by the native skeletal DHPR complex has an inherently low density due to inhibitory interactions within the DHPR and that the ß1a-subunit is critically involved in process.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
The dihydropyridine receptor (DHPR) of skeletal muscle is a heteropentamer composed of 1S, ß1a, 2-1, and 1 subunits. This complex forms a voltage-gated Ca²⁺ channel that is specialized in coupling membrane excitation to the opening or ryanodine receptor type 1 (RyR1) in the sarcoplasmic reticulum (SR). Although excitation-contraction (EC) coupling requires charge movements in the DHPR, the opening of the DHPR Ca²⁺ channel is not essential. Not only is the opening of the Ca²⁺ channel nonessential, but it has long been suspected that the great majority of the skeletal DHPRs do not contribute to the Ca²⁺ current (Schwartz et al., 1985). This has been suggested also by developmental studies showing that skeletal DHPR Ca²⁺ current and charge movements are differentially regulated. The maximal Ca²⁺ conductance and charge movement densities of mouse skeletal myotubes are 140 pS/pF and 7 fC/pF during the late gestation period (Strube et al., 1996, 2000). However, in adult mouse fibers, the Ca²⁺ current density remains at approximately the same level (85 pS/pF, Wang et al., 1999; 160 pS/pF, Jacquemond et al., 2002), but charge movements increase manyfold (40 fC/pF, Wang et al., 1999; 25 fC/pF, Jacquemond et al., 2002). Thus, as the myotube matures, the DHPR complex becomes preferentially engaged in voltage-sensing, and the excess of charge movements is likely to represent a functional adaptation of the complex to an increased demand for EC coupling and force generation.

A critical regulator of DHPR function is the ß-subunit of this complex. Numerous expression studies in heterologous cells have shown that Ca²⁺ channel ß-subunits profoundly affect Ca²⁺ current expression and gating properties of voltage-gated Ca²⁺ channels (Birnbaumer et al., 1998). ß-subunits are cytoplasmic or membrane-associated proteins of 65–75 kDa encoded by four genes. Alternate splicing of the ß1 gene produces at least three isoforms. The ß1a variant is exclusively present in skeletal muscle, whereas the other two (ß1b, ß1c) are expressed in brain, heart and spleen (Powers et al., 1992). Transcripts of the other three genes (ß2, ß3, ß4) were isolated from cardiac and brain cDNA libraries (ß2, Perez-Reyes et al., 1992; ß3, Castellano et al., 1993a; ß4, Castellano et al., 1993b). In terms of sequence content, ß-subunits share two conserved central regions amounting to more than half of the total peptide sequence (domains D2 and D4), a nonconserved linker between the two conserved domains (D3), and nonconserved N- and C-termini (domains D1 and D5, respectively) (Perez-Reyes and Schneider, 1994). The main interaction of ß-subunits with 1 pore subunits is via a conserved 30-residue ß-interaction domain present in domain D4 that binds with high affinity to the cytosolic loop between repeats I and II of the 1 subunit (Pragnell et al., 1994; De Waard et al., 1994). However, secondary interactions with other loops of pore subunits have been documented, especially for the neuronal 1A isoform (Walker et al., 1998). In terms of domain homology, ß-subunits are related to the membrane-associated guanylate kinase superfamily, commonly engaged in membrane signaling. All members of this family have PDZ, SH3, and guanylate kinase homology domains which, in ß-subunits, are present in the conserved D2 and D4 regions (Hanlon et al., 1999).

Structure-function studies have suggested roles for ß-subunits in three distinct areas of DHPR function. First, ß-subunits increase surface expression of Ca²⁺ channels; hence, this protein is vital for trafficking the newly synthesized DHPR out of perinuclear sites (Chien et al., 1995; Neuhuber et al., 1998; Bichet et al., 2000). Second, ß-subunits modulate the kinetics of the L-type Ca²⁺ current (Qin et al., 1998; Wei et al., 2000; Berrou et al., 2001) and increase the coupling between S4 charge movements and pore opening (Neely et al., 1993; Kamp et al., 1996; Olcese et al., 1996). Finally, the ß1a subunit is required for the voltage dependent triggering of Ca²⁺ transients in skeletal muscle, suggesting that ß-participates in coupling charge movements to the opening of RyR1 (Beurg et al., 1999a,b; Sheridan et al., 2002). Studies in HEK cells, aiming to identify the molecular basis of the transport function of ß2a, showed that neither D1 nor D5 was essential for the surface expression of Ca²⁺ channels formed by 1C or for the expression of DHP binding sites accessible from the outside of the cell (Gao et al., 1999). However, mutations in the ß-interaction domain region and the nearby SH3 region abolished surface expression (Chien et al., 1998). Hence, a strong 1/ß interaction is vital for the survival of the newly synthesized Ca²⁺ channel. In terms of the functional identity of other domains present in the ß-subunit, the N-terminus in domain D1 is a critical determinant of the rate and voltage dependence of activation of Ca²⁺ channels formed by the 1E isoform (Olcese et al., 1994). A second region in domain D3 selectively affected inactivation independent of ß-effects on activation (Qin et al., 1996). Furthermore, the C-terminus region of ß1a (domain D5) is a critical determinant of the EC coupling function of the DHPR in skeletal myotubes (Beurg et al., 1999a,b; Sheridan et al., 2002).

A second critical regulator of DHPR function is RyR1 itself. Studies in RyR1 KO myotubes have shown that approximately half of the DHPR charge movements and DHP binding sites are lost when RyR1 is eliminated. However, the Ca²⁺ current density is depressed much more severely, close to10-fold (Nakai et al., 1996; Buck et al., 1997; Avila and Dirksen, 2000). Expression of the DHPR Ca²⁺ current has been suggested to be under the specific control of RyR1 by a so-called retrograde signal, from RyR1 to the DHPR (Nakai et al., 1998). The nature of this retrograde signal is probably structural in origin because full Ca²⁺ current recovery was observed in RyR KO myotubes expressing RyR1 mutants strongly deficient in voltage-dependent EC coupling (Avila et al., 2001). However, full charge movement recovery in the RyR1 KO myotube did not occur with the EC coupling deficient RyR1 mutant, requiring instead an entirely functional RyR1 (Avila et al., 2001). Thus, the contribution of RyR1 to DHPR expression is complex and presumably involves modulatory effects via protein-protein interactions as well as Ca²⁺ signaling mediated by RyR1. In contrast to the strict requirements for RyR1 in the skeletal myotube, studies in amphibian oocytes have shown that skeletal Ca²⁺ current expression can be achieved by a specific combination of skeletal (1S, 2, 1) and neuronal (ß1b) subunits (Ren and Hall, 1997; Morrill and Cannon, 2000). The reasons why the DHPR Ca²⁺ current would require a retrograde signal from RyR1 in the skeletal myotube but this signal would be bypassed in the heterologous oocyte system are entirely unclear at this point. This discrepancy suggests that in the skeletal myotube, Ca²⁺ current regulation is multifactorial, and perhaps it represents a balance of positive and negative interactions within the DHPR and between the DHPR and other muscle proteins.

In this study, we investigated the charge movement and Ca²⁺ current characteristics donated to this complex by ß1a, the endogenous variant known to form part of the skeletal DHPR (Ruth et al., 1989), and by ß2a (Perez-Reyes et al., 1992; Qin et al., 1998), which is a variant expressed in the heart and brain but not in skeletal muscle (Perez-Reyes and Schneider, 1994). Studies were conducted in RyR1 KO, ß1 KO, and double KO myotubes generated by interbreeding mice heterozygous for the ß1 KO and RyR1 KO alleles. The data indicate that in the absence of RyR1, ß2a had a preferential ability to recover a DHPR that expressed Ca²⁺ currents at a high density and charge movements at a low density. Conversely, ß1a had a preferential ability to recover a DHPR that expressed charge movements at a high density and Ca²⁺ currents at a low density. Thus, the ß-subunit of the skeletal DHPR appears to be intimately involved in the cellular mechanisms that differentially regulate charge movements and Ca²⁺ current expression in the skeletal myotube. Part of this work has been previously published in abstract form (Ahern et al., 2002).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Identification of genotypes
We used PCR to screen for the wild-type (WT) and knock-out (KO) alleles of the DHPR ß1 and RyR1 genes in mice with targeted disruptions of these genes (Gregg et al., 1996; Nakai et al., 1996). Tail samples from E18 fetuses were digested with Proteinase K (Sigma, St. Louis, MO), and the DNA was then isolated following the Puregene animal tissue protocol (Gentra Systems, Minneapolis, MN). The PCR reactions for each sample were composed of 11.7 µL distilled water, 1 µL of each 20 µM primer, 3.2 µL of 1.25 mM dNTPs (Stratagene, Cedar Creek, TX), 2 µL 10x PCR buffer (Qiagen, Valencia, CA), 12 µL Taq polymerase (Qiagen, Valencia, CA), and 1 µL of the DNA solution (100 µg/mL). PCR primers 5' gga ctg gca aga gga ccg gag 3' and 5' gga agc cag ggc tgc agg tga gc 3' were used to amplify a 400 bp fragment of the WT RyR1 allele. PCR primers 5' gga ctg gca aga gga ccg gag 3' and 5' cct gaa gaa cga gat cag cag cct ctg ttc c 3' were used to amplify a 300 bp fragment of the KO RyR1 allele. The following conditions applied for the PCR of the RyR1 alleles: 1) 5 min and 30 s at 94°C, 2) 1 min at 94°C, 3) 1 min at 55°C, 4) 1 min at 72°C, 5) cycle through steps 2–4 for 30 times, and 6) 10 min at 72°C. PCR primers 5' cta gga gaa tgg atg gta gat gg 3' and 5' aca ccc cct gcc agt ggt aag agc 3' were used to amplify a 250 bp fragment of the WT ß1 allele. PCR primers 5' aca ccc cct gcc agt ggt aag agc 3' and 5' aca ata gca ggc atg ctg ggg atg 3' were used to amplify a 197 bp fragment of the KO ß1 allele. The following conditions apply for the PCR of the DHPR ß1 alleles: 1) 2 min at 94°, 2) 30 s at 94°C, 3) 45 s at 55°C, 4) 1 min at 72°C, 5) cycle through steps 2–4 for 30 times, and 6) 10 min at 72°C.

Myotube primary cell cultures
Cell cultures of myotubes were prepared from hind limbs of E18 mice as described (Beurg et al., 1997). Myoblasts were isolated by enzymatic digestion with 0.125% (w/v) trypsin and 0.05% (w/v) pancreatin. After centrifugation, mononucleated cells were resuspended in plating medium containing 78% Dulbeccos's modified Eagle's medium with low glucose (DMEM), 10% horse serum, 10% fetal bovine serum (FBS), and 2% chicken serum extract. Cells were plated on plastic culture dishes coated with gelatin at a density of 1 x 10⁴ cells per dish. Cultures were grown at 37°C in 8% CO₂ gas. After the fusion of myoblasts (5–7 days), the medium was replaced with FBS-free medium, and CO₂ was decreased to 5%.

cDNA transfection
Cell transfection was performed during myoblast fusion with the polyamine LT1 (Panvera). Cells were exposed for 2–3 h to a transfection solution containing LT1 and cDNA in a 5:1 µl/µg ratio. All cDNAs of interest, with the exceptions of RyR1 and CD8-ß2a C3,4S, were subcloned in the mammalian expression vector pSG5 (Stratagene, La Jolla, CA). The RyR1 cDNA was subcloned in the pCl-neo expression vector (Promega, Madison, WI) as described (Nakai et al., 1996). The CD8-ß2a C3,4S construct is described below. The expression plasmid was cotransfected with a plasmid encoding the T-cell surface glycoprotein CD8A used as a transfection marker. Transfected cells were identified with micron-size polystyrene beads coated with a monoclonal antibody against an external CD8A epitope (Dynal, Oslo, Norway). The efficiency of cotransfection of the marker and the cDNA of interest was 80%.

Full-length cDNAs
All cDNAs were N-terminus tagged by in-frame fusion of the first 11 amino acids of the phage T7 gene 10 protein in the pSG5 vector. GenBank accession numbers for the full-length sequences utilized in the present study were as follows: mouse ß1a residues 1–524 (NM_031173), mouse ß1b residues 1–596 (AY094173), mouse ß1c residues 1–480 (AY094172), rat ß2a residues 1–604 (M80545), rat ß3b residues 1–485 (M88751), and rat ß4 residues 1–519 (L02315). All cDNA constructs were sequenced at least twice at a campus facility.

ß1a deletion constructs
Two oligonucleotide primers were designed to encompass the region of interest. Each primer had 20–25 bases identical to the original sequence and an additional 10–15 bases that resulted in an amplified product with an AgeI site at the 5' end and a stop codon and NotI restriction site at the 3' end. The PCR products were subcloned into the pCR-Blunt vector (Invitrogen, Carlsbad, CA), excised by digestion with AgeI and NotI, and cloned into pSG5.

ß1a1–57
PCR primers 5' gcat gac cgg tgg aca gca aat ggg tgg ctc agc aga gtc cta cac 3' and 5' gcg gcc gct agc tac cta cat ggc gtg ctc ctg agg 3' were used to generate a cDNA that contains amino acids 58–524 of the ß1a cDNA. This cDNA was fused in frame to the first 11 amino acids of the phage T7 gene 10 protein in the pSG5 vector.

ß1a490–524
PCR primers 5' gca tga ccg gtg gac agc aaa tgg gta tgg tcc aga aga gcg gca tgt ccc ggg gcc 3' and 5' ggg gcg gcc gct cac tgg agg ttg gag acg ggg gca 3' were used to generate a cDNA that contains amino acids 1–489 of the ß1a cDNA. This cDNA was fused in frame to the first 11 amino acids of the phage T7 gene 10 protein in the pSG5 vector.

Cysteine substitution constructs
ß1a Q3C, K4C
Forward primer 5' acc ggt gga aca gca aat ggg tat ggt ctg ctg cag ggg cat g 3' and reverse primer 5' att gta gcc aac att tgt ccg aac agc 3' were used to introduce nucleotides for cysteines at residue positions three and four in the ß1a template. Primers (10 pmol each), 20 pmol dNTPs, 20 ng T7-ß1a-pSG5 template, and 0.5 µl Taq polymerase (Promega, Madison, WI) were combined and cycled at 95°C for 5 min then 95°C for 2 min, 55°C for 1 min, and 72°C for 1 min for 30 cycles ending with 10 min at 72°C. The PCR product was ligated into pCR 2.1 TA vector (Stratagene, Carlsbad, CA). This cDNA was then inserted into T7-ß1a-pSG5 via unique AgeI and XhoI sites.

ß2a C3,4S
Forward primer 5' acc ggt gga cag caa atg gta tgc agt cct ccg ggc 3' and reverse primer 5' cat atc tgt gac ctc ata cc 3' were used to introduce nucleotides for serines at residue positions three and four in the ß2a template. Primers (10 pmol each), 20 pmol dNTPs, 20 ng T7-ß2a-pSG5 template, and 0.5 µl Taq polymerase (Promega, Madison, WI) were combined and cycled at 95°C for 5 min then 95°C for 2 min, 55°C for 1 min and 72°C for 1 min for 30 cycles finishing with 10 min at 72°C. The PCR product was cloned into pCR 2.1 TA vector (Stratagene, Carlsbad, CA). This cDNA was then inserted into T7-ß2a-pSG5 via unique AgeI and Acc65I sites.

Chimeric ß2aD1–3ß1aD4,5 (ß2a 1–287/ß1a 325–524)
This construct contains amino acids 1–287 of ß2a fused to the N-terminus of amino acids 325–524 of ß1a and was made by two rounds of PCR. Primers 5' gca tga ccg gtg gac agc aaa tgg gta tgc agt gct gcg ggc tgg ta 3' (A) and 5' gag cgt ttg gcc agg gag atg tca gca 3' (B) were used to PCR the 5' end of the full-length ß1a cDNA. Primers 5' tcc ctg gcc aaa cgc tcc gtc ctc aac 3' (C) and 5' gcg gcc gct agc tac cta cat ggc gtg ctc ctg agg 3' (D) were used to PCR the 3' end of the full-length ß1a cDNA. The primers were designed to produce a 17 bp overlap of identical sequence. The two PCR products were electrophoresed on agarose gels, excised from the gel, and eluted using GenElute columns (Supelco, Bellefonte, PA). The two PCR products were mixed in an equimolar ratio, denatured, allowed to reanneal, and used in a PCR reaction to amplify the chimeric fragments with primers A and D. This cDNA was fused in frame to the first 11 amino acids of the phage T7 gene 10 protein in the pSG5 vector.

Chimeric ß2aD1ß1aD2–5 (ß2a 1–16/ß1a 58–524)
This construct contains amino acids 1–16 of ß2a cDNA fused to the N-terminus of amino acids 58–524 of ß1a cDNA and was made by one round of PCR. Forward primer 5' gga tcc atg cag tgc tgc ggg ctg gta cat cgc cgg cga gta cgg gtc cgc cag ggc tca gc 3' encodes a portion of the N-terminal T7 tag, the first 16 residues of ß2a and ß1a residues 58–67. This "overhang" primer was combined with reverse primer 5' att gta gcc aac att tgt ccg aac agc 3' to produce the chimeric construct containing domain D1 of ß2a fused to domain D2 to D5 of ß1a. Primers (10 pmol each), 20 pmol dNTPs, 20 ng T7-pSG5-ß1a template, and 0.5 µl Taq polymerase (Promega, Madison, WI) were combined and cycled at 95°C for 5 min then 95°C for 2 min, 55°C for 1 min and 72°C for 1 min for 30 cycles ending with 10 min at 72°C. The PCR product was cloned into pCR 2.1 TA vector (Stratagene). This cDNA was then inserted into the T7-ß1a-pSG5 vector via unique AgeI and XhoI sites.

Chimeric ß1aD1–D3ß2aD4,5 (ß1a 1–325/ß2a 287–604)
This construct contains amino acids 1–325 of ß1a cDNA fused to the N-terminus of amino acids 287–604 of ß2a cDNA and was made by two rounds of PCR. In the first round, the ß1a cDNA fragment was amplified by the forward primer 5' caa gac tag cgt gag cag tgt cac c 3' (A) encoding residues 240–247 of ß1a and the reverse primer 5' gtg ttg gat ctt tct atg acg gag cgt ttg 3' (B), which encodes residues 321–325 of ß1a and residues 287–292 of ß2a. The ß2a cDNA fragment was amplified by the forward primer 5' ata gaa aga tcc aac aca agg tcg agc 3' (C) encoding residues 287–295 of ß2a and the reverse primer 5' aag ctg caa taa aca agt tct gc 3' (D) which is homologous to the pSG5-T7 vector sequence. The first two PCR products were mixed after purification with QIAquick PCR purification columns (QIAGEN, Valencia, CA), and were annealed and extended by Taq polymerase in the absence of primers (95°C for 2 min, then 95°C for 1 min, 43°C for 1 min, and 72°C for 1 min for 5 cycles). Afterward, primers A and D were added to the PCR mixture and the reaction was continued at 95°C for 1 min, 65°C for 1 min, and then 72°C for 1 min for 25 cycles ending with 10 min at 72°C. The final PCR product was subcloned into pCR 2.1 TA vector (Stratagene). This cDNA was then inserted into the T7-ß1a-pSG5 vector via unique PmlI and NotI sites.

Chimeric ß2aC3,4SD1–3ß1aD4,5 (ß2a C3,4S 1–287/ß1a 325–524)
This construct contains amino acids 1–287 of ß2a fused to the N-terminus of amino acids 325–524 of ß1a with cysteines substituted for serines at positions three and four of ß2a. Forward primer 5' acc ggt gga cag caa atg gta tgc agt cct ccg ggc 3'and reverse primer 5' cat atc tgt gac ctc ata cc 3' were used to introduce nucleotides for serines at residue positions three and four in the chimeric ß2a 1–287/ß1a 325–524 template. Primers (10 pmol each), 20 pmol dNTPs, 20 ng T7-pSG5-ß2a template, and 0.5 µl Taq polymerase (Promega, Madison, WI) were combined and cycled at 95°C for 5 min then 95°C for 2 min, 55°C for 1 min and 72°C for 1 min for 30 cycles finishing with 10 min at 72°C. The PCR product was cloned into pCR 2.1 TA vector (Stratagene). This cDNA was then inserted into T7-ß2a 1–287/ß1a 325–524-pSG5 vector via unique AgeI and Acc65I sites.

Chimeric CD8-ß2a C3,4S
This construct was described previously (Restituo et al., 2000). Residues 1–209 of the human CD8A surface glycoprotein (accession number NM_001768) were fused in-frame to the N-terminus of ß2a C3S,C4S in expression vector pcDNA3 (Invitrogen, Carlsbad, CA). The N-terminus of CD8 has an external epitope recognized by anti-CD8 monoclonal antibody 5C2 adsorbed on polystyrene Dynabeads M-450 (Dynal).

Sequence alignment
Alignment of mouse ß1a (NM_031173) and rat ß2a (M80545) peptide sequences was performed with DNASTAR (Madison, WI) using the Jotun Hein method and an identity residue weight table. Other alignment methods and residue weight tables also produced similar results. Based on this alignment, domain D1 corresponds to residues 1–57 of ß1a and residues 1–16 of ß2a with a similarity of 41.2%; D2 corresponds to residues 58–198 of ß1a and residues 17–157 of ß2a with a similarity of 78.2%; D3 corresponds to residues 199–253 of ß1a and residues 158–205 of ß2a with a similarity of 36.7%; D4 corresponds to residues 254–477 of ß1a and residues 206–429 of ß2a with a similarity of 90.6%; and D5 corresponds to residues 478–524 of ß1a and residues 430–604 of ß2a with a similarity of 21.7%.

Whole-cell voltage-clamp
Cells were voltage-clamped 3–5 days after transfection. Transfected cells revealed by CD8 beads were voltage-clamped with an Axopatch 200B amplifier (Axon Instruments, Foster City, CA) and a Digidata 1200 (Axon Instruments) pulse generator and digitizer. Linear capacitance, leak currents, and effective series resistance were compensated with the amplifier circuit. The charge movement protocol included a long pre-pulse to eliminate immobilization-sensitive gating currents. Voltage was stepped from a holding potential of -80 mV to -30 mV for 700 ms, then to -50 mV for 5 ms, then to the test potential for 50 ms, then to -50 mV for 30 ms, then to -80 mV. Test potentials were applied in decreasing order every 10 mV from +100 or +110 mV to -70 mV. The intertest pulse period was 10 s. Subtraction of the linear charge was done online with a P/4 procedure. The P/4 pulses were delivered immediately before the protocol from -80 mV in the negative direction. The P/4 pulses adequately subtracted the linear component of the charging current, in the range of -80 to -120 mV. In this range, the membrane capacity varied linearly with voltage, within a 0.5% error.

Solutions
The external solution in all cases was (in mM) 130 TEA methanesulfonate, 10 CaCl₂, 1 MgCl₂, 10 HEPES titrated with TEA(OH) to pH 7.4. For Ca²⁺ currents, the pipette solution was (in mM) 140 Cs⁺-aspartate, 5 MgCl₂, 5 EGTA, and 10 MOPS-CsOH, pH 7.2. For charge movements, the pipette solution was (in mM) 120 NMG (N-methyl glucamine)-glutamate, 10 HEPES-NMG, 10 EGTA-NMG, pH 7.3 (Ahern et al., 2001b). For charge movements, the external solution was supplemented with 0.5 mM CdCl₂, 0.5 mM LaCl₃, and 0.05 mM TTX to block residual Na⁺ current. For Ca²⁺ transients, the pipette solution was (in mM) 140 Cs⁺-aspartate, 5 MgCl₂, 0.1 EGTA, and 10 MOPS-CsOH, pH 7.2.

Confocal fluorescence microscopy
Confocal line scan measurements were performed as described previously (Conklin et al., 1999). Cells were loaded with 4 µM fluo-4 acetoxymethyl (AM) ester (Molecular Probes, Eugene, OR) for 60 min at room temperature. All experiments were performed at room temperature. Cells were viewed with an inverted Olympus microscope with a 20x objective (N.A. = 0.4) and an Olympus Fluoview confocal attachment (Melville, NY). A 488 nm spectrum line for fluo-4 excitation was provided by a 5 mW Argon laser attenuated to 20% with neutral density filters. Line scans consisted of 1,000 lines, each of 512 pixels, acquired at a rate of 2.05 ms per line. Line scans were synchronized to start 100 ms before the onset of the depolarization. The time course of the space-averaged fluorescence intensity change was estimated as follows. The pixel intensity was transformed into arbitrary units, and the mean intensity was obtained by averaging pixels covering the cell exclusively. The mean resting fluorescence intensity, Fo, corresponds to the mean intensity of each line in the line scan averaged over the first 100 ms before stimulation. The change in mean intensity above rest, F, was obtained by subtraction of Fo from the mean intensity of each line in the line scan. F for each line in a line scan was divided by Fo and the ratio F/Fo was plotted as a function of time. To construct peak Ca²⁺ fluorescence versus voltage curves, we used the highest F/Fo line value after the onset of the pulse and up to the termination of the pulse. Image analysis was performed with NIH Image software (National Institutes of Health, Bethesda, MD). To obtain reliable Ca²⁺ transient versus voltage curves, seven step depolarizations of 50 ms were applied in descending order (from +90 mV to -30 mV) from a holding potential of -40 mV. Between each depolarization, the cell was maintained at the resting potential for 30 s to permit recovery of the resting fluorescence.

Curve fitting
The voltage dependence of charge movements (Q), Ca²⁺ conductance (G), and peak fluorescence F/Fo was fitted according to a Boltzmann distribution (Eq. 1), namely A = Amax/(1+exp(-(V-V_1/2)/k), where Amax is Qmax, Gmax, or F/Fo max; V_1/2 is the potential at which A = Amax/2; and k is the slope factor. The statistic of Boltzmann parameters (Amax, V_1/2, k) were obtained by fitting curves to individual cells and are shown in Tables 1 and 2. The figures show the fit of the mean Amax of all cells with the number of cells shown in Tables 1 and 2. The statistical analyses were performed with Analyse-it software (Analyse-It Software, Leeds, UK).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

View larger version (95K): [in this window] [in a new window]   FIGURE 1  Genotype of ß1/RyR1 KO mice obtained by PCR. DNA fragments amplified with genotype-specific primers are shown on two separate 3% agarose gels. Each gel shows two sets of PCR reactions, each for a specific allele (top: RyR1 KO, RyR1 WT; bottom: ß1 KO, ß1 WT). Indicated on the left are three size markers (500, 200, and 100 bp) of a total of six markers run in each gel. Indicated by arrows are the expected sizes of the amplified DNA for each reaction. Templates used were total DNA of three paralytic littermates (lanes 1–3), a double heterozygous parent (lane 4), and a control reaction without template (lane 5).

View larger version (23K): [in this window] [in a new window]   FIGURE 2  DHPR Ca2+ currents in ß1/RyR1 KO, RyR1 KO, and ß1 KO myotubes. Ca2+ currents are shown in response to 500-ms voltage steps to -10, +10, and +30 mV from a holding potential of -40 mV. Nontransfected (NT), ß1a-expressing, or ß2a-expressing myotubes are shown in the top, middle and bottom rows, respectively. Scale bars are 1 nA and 200 ms. The graphs show the voltage dependence of the Ca2+ conductance calculated from the maximal current during the pulse and the reversal potential obtained by extrapolation. Symbols show the mean Ca2+ conductance density (± SE) for the number of cells indicated in Table 1. The mean was fitted according to Eq. 1. The parameters of the fit with Gmax in pS/pF, V1/2 in mV, and k in mV are as indicated below. For ß1/RyR1 KO myotubes 29.8, 14.8, 7.4 for ß1a-transfected; and 114.8, 9.7, 7.5 for ß2a-transfected. For RyR1 KO myotubes 37.8, 15.0, 8.6 for NT; 35.8, 7.1, 7.1 for ß1a-transfected; and 196.7, 10.8, 5.1 for ß2a-transfected. For ß1 KO myotubes 156.6, -0.3, 4.9 for ß1a-transfected; and 174.8, 7.5, 5.3 for ß2a-transfected.

A higher Ca²⁺ current density recovered by ß2a compared to ß1a could reflect a better ability of ß2a to traffic the DHPR to the cell surface. To determine if this was the case, we used gating currents to gauge the membrane density of DHPR voltage sensors. We have previously shown that a pore-forming DHPR generates as much gating current as a non pore-forming DHPR (Ahern et al., 2001a). Thus, the technique is impervious to the functional state of the Ca²⁺ channel, which is an important consideration in the present case. Fig. 3 shows intramembrane currents produced by the movement of charges in the expressed DHPR in response to 4 of 17 step voltages delivered to each cell (-30, +10, +50, and +90 mV) from a holding potential of -80 mV. The protocol included a 700-ms pre-test pulse depolarization to -30 mV to remove immobilization sensitive components of the gating current unrelated to the DHPR and pre-test P/4 pulses to eliminate linear components unrelated to voltage sensors. We found a good correspondence between the magnitude of the ON and OFF components, as would be expected if charge movements were due to reversible displacements of intramembrane charges. The graphs at the bottom of each column in Fig. 3 show charge versus voltage relationships for nontransfected and ß1a- and ß2a-expressing myotubes. The immobilization-resistant charge was obtained by integration of the OFF component which, in our hands, was less contaminated by ionic current than the ON component. The main contaminant of the ON component was a nonlinear outward current, presumably from a K⁺ channel, that was not always blocked by the pipette solution. In all cases, the OFF charge increased in a sigmoidal fashion starting at approximately -10 mV and saturated at potentials more positive than +60 mV, and data were adequately fit by a Boltzmann equation (Eq. 1) shown by the lines. In the ß1/RyR1 KO myotube (Fig. 3, left column), we detected a small background charge with a maximal density of 1 fC/pF. Overexpression in these cells of ß1a or ß2a increased the maximal charge density threefold and twofold, respectively, and this difference was significant in each case (see Table 1). Thus, contrary to expectations based on Ca²⁺ current densities, the density of DHPR voltage sensors was lower in ß2a-expressing than in ß1a-expressing double KO myotubes. Overexpression of ß1a in the double KO myotube reconstituted quantitatively the density of DHPR voltage sensors present in the nontransfected RyR1 KO myotube. This can be seen by comparing the charge versus voltage curves at the bottom of Fig. 3 (left plot open symbols versus center plot gray symbols) and the statistic of the Boltzmann parameters presented in Table 1. The Qmax of ß1a-expressing ß1/RyR1 KO myotubes was 3.3 ± 0.4 fC/pF, whereas the Qmax of nontransfected RyR1 KO myotubes was 3.9 ± 0.4 fC/pF, with the latter result in agreement with previous determinations (Avila and Dirksen, 2000). Interestingly, neither overexpression of ß1a nor ß2a rescued additional charge in the RyR1 KO myotube. ß2a caused a slight reduction in Qmax relative to the nontransfected background, although this difference was not significant. The same pattern of charge recovery observed in the double KO was repeated in the ß1 KO myotube (Fig. 3, right column). The maximal charge density in the ß1a-expressing ß1 KO myotube was higher than in ß2a-expressing myotubes. However, due to the background charge present in nontransfected cells, the charge expressed by ß1a, but not that by ß2a, was significantly different from background (see Table 1). The Qmax of nontransfected ß1 KO myotubes was twofold higher than the Qmax of the ß1/RyR1 KO myotube or that of the 1S-null dysgenic myotube (Ahern et al., 2001a). Such excess of charges is likely to arise from a low density of ß1-less DHPR complexes transported to the surface, perhaps by residual levels of other beta isoforms present in the skeletal myotube. In summary, the intramembrane charge movement analyses did not provide evidence for a differential ability of ß2a to promote transport and targeting of the DHPR to the cell surface. On the contrary, ß2a was less effective than ß1a in all three myotube genotypes analyzed.

View larger version (29K): [in this window] [in a new window]   FIGURE 3  DHPR charge movements in ß1/RyR1 KO, RyR1 KO, and ß1 KO myotubes. Intramembrane currents are shown in response to 50-ms voltage steps to -30, +10, +50, and +90 mV from a holding potential of -80 mV. Nontransfected (NT), ß1a-expressing, or ß2a-expressing myotubes are shown in the top, middle, and bottom rows, respectively. Scale bars are 0.5 nA and 25 ms. The graphs show the voltage dependence of the intramembrane charge obtained by integration of the OFF current. Symbols show the mean nonlinear charge density (± SE) for the number of cells indicated in Table 1. The lines are a fit of the mean according to Eq. 1. The parameters of the fit with Qmax in fC/pF, V1/2 in mV, and k in mV are: for ß1/RyR1 KO myotubes 1.3, 18.5, 16.6 for NT; 3.1, 15.1, 17.3 for ß1a-transfected; and 2.2, 16.7, 17.2 for ß2a-transfected. For RyR1 KO myotubes 3.8, 16.2, 17.8 for NT; 3.4, 16.3, 23.4 for ß1a-transfected; and 2.5, 12.1, 19.4 for ß2a-transfected. For ß1 KO myotubes 2.0, 1.6, 19.2 for NT; 4.6, 25.7, 19.2 for ß1a-transfected; and 3.4, 15.6, 19.5 for ß2a-transfected.

View larger version (16K): [in this window] [in a new window]   FIGURE 4  Kinetics of activation of Ca2+ currents in ß1 KO myotubes expressing ß1a or ß2a. A, B) Ca2+ currents (gray line) were normalized according to the maximal current during the pulse. They were obtained in ß1 KO myotubes transfected with ß1a (top) or ß2a (bottom) in response to a 500-ms voltage step to +30 mV from a holding potential of -40 mV. Scale bar is 100 ms. To fit the current during the pulse, the first 10 ms after the onset of the pulse was omitted. A) shows a monoexponential fit and B) shows a biexponential fit of the same data, with the fit indicated by the black line. Time constants of the monoexponential fit are 58.4 ms for ß1a and 103.4 ms for ß2a. Time constants of the biexponential fit are 7, 62.5 ms for ß1a and 16.9, 119.2 ms for ß2a. Relative contributions of the fast and slow components are 0.11, 0.89 for ß1a and 0.15, 0.85 for ß2a. Chi-square tests of the fit were as follows: 2 = 0.026 for monexponential and 2 = 0.00024 for biexponential fits of ß1a; 2 = 0.028 for monoexponential and 2 = 0.0006 for biexponential fits of ß2a. C) Voltage dependence of the single activation time constant obtained from the monoexponential fit of Ca2+ currents. Unpaired t-tests revealed no significant differences in a comparison of ß1a (empty symbols) versus ß2a (filled symbols) at any voltage. D) Voltage dependence of the activation time constants obtained from the biexponential fit of the same data. Unpaired t-test revealed significance differences (p < 0.05) for slow at -5, 0, +5, +10, and +15 mV, and fast at +15, +20, +25, and +30 mV.

View larger version (22K): [in this window] [in a new window]   FIGURE 5  Kinetics of activation of Ca2+ currents expressed by ß2a in the absence and presence of RyR1. A) Normalized Ca2+ currents (gray) are shown in response to a 500-ms depolarization to +30 mV from a holding potential of -40 mV. Scale bar is 100 ms. The first 10 ms after the onset of the pulse was omitted, and the remainder of the pulse current was fit as a biexponential in all cases (black line). The top row shows the fit of the pulse current in normal (WT), RyR1 KO, and RyR1-expressing RyR1 KO myotubes. The bottom row shows the fit in ß2a-expressing ß1/RyR1 KO, RyR1 KO, and ß1 KO myotubes. fast and slow are 8.6, 74.5; 4.3, 34.3; 6.1, 70.2; 16.3, 112.9; 16.3, 110.3; 16.9, 119.2 ms, respectively, from left to right with top row first. Relative contributions of fast and slow components are 0.24, 0.76; 0.26, 0.74; 0.28, 0.72; 0.16, 0.84; 0.20, 0.80; 0.15, 0.85, respectively, in the same order. B) The mean (+1 SE) fast and slow at +30 mV is shown for the genotypes analyzed above. The last three entries correspond to myotubes expressing ß2a. The number of fitted Ca2+ currents, each from a separate myotube, included in the averages was 7 (WT), 10 (RyR1 KO), 10 (RyR1 KO + RyR1), 5 (ß1/RyR1 KO + ß2a), 10 (RyR1 KO + ß2a), and 8 (ß1 KO + ß2a). One-way ANOVA showed no significant difference for fast (p = 0.75) or slow (p = 0.27) of ß2a-expressing myotubes. All other entries were significantly different from ß2a-expressing myotubes (p < 0.001). C) The mean (+1 SE) contribution of the fast and slow components of the biexponential fit is shown for all genotypes analyzed. One-way ANOVA revealed no significant differences between fast and slow components analyzed separately.

View larger version (19K): [in this window] [in a new window]   FIGURE 6  Ca2+ conductance recovered by DHPR ß variants in RyR1 KO myotubes. The Ca2+ conductance of RyR1 KO myotubes transfected with the indicated ß-construct was fit to Eq. 1 for the number of cells indicated in parenthesis. The ß2a-ß1a chimera was ß2a 1–287/ß1a 325–524. The ß1a-ß2a chimera was ß1a 1–325/ß2a 287–604. Gmax in pS/pF, V1/2 in mV, and k in mV were: 37.2 ± 5.2, 12.4 ± 2.4, 6.3 ± 0.3 for NT; 35.6 ± 5.6, 9.9 ± 3.7, 5.4 ± 0.3 for ß1a; 41.7 ± 3.7, 18.1 ± 2.3, 6.8 ± 1.1 for ß1b; 34.8 ± 6.4, 23 ± 2.4, 7.3 ± 1.7 for ß1c; 195 ± 16, 10.8 ± 1.3, 4.9 ± 0.2 for ß2a; 39.7 ± 6.5, 19.7 ± 2.8, 5.8 ± 0.5 for ß3; 56.8 ± 10.6, 26.5 ± 2.6, 5.8 ± 0.5 for ß4; 36.3 ± 11.5, 14.6 ± 2.0, 6.3 ± 0.6 for ß1a 1–57; 41.3 ± 7.6, 17.9 ± 1.1, 6.3 ± 0.3 for ß1a 490–524; 173 ± 14.0, 9.3 ± 2.4, 4.3 ± 0.6 for ß2a-ß1a; and 38.6 ± 8.2, 10.6 ± 3.4, 5.9 ± 2.2 for ß1a-ß2a. Unpaired t-test showed significant differences (p < 0.001) relative to nontransfected for the maximal Ca2+ conductance recovered by ß2a and ß2a-ß1a.

View larger version (53K): [in this window] [in a new window]   FIGURE 7  Participation of a double cysteine motif of ß2a in Ca2+ current expression in RyR1 KO myotubes. A) Comparison of residues 1 through 14 of mouse ß1a, rat ß2a, rat ß3b, and rat ß4. Nonconserved cysteines at positions three and four of ß2a are underlined. B) Ca2+ current expression by a ß2a variant lacking cysteines at positions three and four (ß2a C3,4S) and by a membrane-anchored variant CD8-ß2a C3,4S in RyR1 KO myotubes. Ca2+ currents were elicited by 500-ms depolarizations from a holding potential of -40 mV to -10, +10, and +30 mV. Scale bars are 2 nA and 100 ms. C) Wide-field view of RyR1 KO myotubes expressing CD8-ß2a C3,4S. Surface expression of the construct is revealed by anti-CD8 beads added to the external solution. Myotubes were transfected with the CD8-ß2a C3,4S cDNA exclusively. Asterisk marks nontransfected myotubes in the same field. Bar is 50 microns and bead diameter is 4.5 microns. Diagrams depict membrane-anchoring of wild-type ß2a and CD8-ß2a C3,4S according to previous studies (Chien et al., 1996; Qin et al., 1998; Restituto et al., 2000). D) Voltage dependence of the Ca2+ conductance in RyR1 KO myotubes. The line without data corresponds to the voltage dependence of the Ca2+ conductance of ß2a-expressing RyR1 KO myotubes from Fig. 2. Symbols show the mean conductance (± SE) for the number of cells indicated in Table 2. The lines are a fit of the mean conductance according to Eq. 1. Parameters of the fit with Gmax in pS/pF, V1/2 in mV, and k in mV are 56, 16.8, and 6.6 for ß2a C3,4S (open diamonds); and 160, 21.7, and 6.8 for CD8-ß2a C3,4S (filled diamonds).

View larger version (17K): [in this window] [in a new window]   FIGURE 8  Participation of domains D1–D3 of ß2a in Ca2+ current expression in RyR1 KO myotubes. A) Ca2+ currents are shown in RyR1 KO myotubes expressing the indicated construct. The diagrams depict domains D1–D5 with ß1a seque