Biophysical Journal 85:3739-3757 (2003)
© 2003 The Biophysical Society
Ca2+-Dependent Excitation-Contraction Coupling Triggered by the Heterologous Cardiac/Brain DHPR ß2a-Subunit in Skeletal Myotubes
David C. Sheridan,
Leah Carbonneau,
Chris A. Ahern,
Priya Nataraj and
Roberto Coronado
Department of Physiology, University of Wisconsin, School of Medicine, Madison, Wisconsin 53706 USA
Correspondence: Address reprint requests to Roberto Coronado, Dept. of Physiology, University of Wisconsin, 1300 University Ave., Madison, WI 53706. Tel.: 608-263-7487; Fax: 608-265-5512; E-mail: coronado{at}physiology.wisc.edu.
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ABSTRACT
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Molecular determinants essential for skeletal-type excitation-contraction (EC) coupling have been described in the cytosolic loops of the dihydropyridine receptor (DHPR) 
1S pore subunit and in the carboxyl terminus of the skeletal-specific DHPR ß1a-subunit. It is unknown whether EC coupling domains present in the ß-subunit influence those present in the pore subunit or if they act independent of each other. To address this question, we investigated the EC coupling signal that is generated when the endogenous DHPR pore subunit 1S is paired with the heterologous heart/brain DHPR ß2a-subunit. Studies were conducted in primary cultured myotubes from ß1 knockout (KO), ryanodine receptor type 1 (RyR1) KO, ryanodine receptor type 3 (RyR3) KO, and double RyR1/RyR3 KO mice under voltage clamp with simultaneous monitoring of confocal fluo-4 fluorescence. The ß2a-mediated Ca2+ current recovered in ß1 KO myotubes lacking the endogenous DHPR ß1a-subunit verified formation of the 1S/ß1a pair. In myotube genotypes which express no or low-density L-type Ca2+ currents, namely ß1 KO and RyR1 KO, ß2a overexpression recovered a wild-type density of nifedipine-sensitive Ca2+ currents with a slow activation kinetics typical of skeletal myotubes. Concurrent with Ca2+ current recovery, there was a drastic reduction of voltage–dependent, skeletal-type EC coupling and emergence of Ca2+ transients triggered by the Ca2+ current. A comparison of ß2a overexpression in RyR3 KO, RyR1 KO, and double RyR1/RyR3 KO myotubes concluded that both RyR1 and RyR3 isoforms participated in Ca2+-dependent Ca2+ release triggered by the ß2a-subunit. In ß1 KO and RyR1 KO myotubes, the Ca2+-dependent EC coupling promoted by ß2a overexpression had the following characteristics: 1), L-type Ca2+ currents had a wild-type density; 2), Ca2+ transients activated much slower than controls overexpressing ß1a, and the rate of fluorescence increase was consistent with the activation kinetics of the Ca2+ current; 3), the voltage dependence of the Ca2+ transient was bell-shaped and the maximum was centered at 
+30 mV, consistent with the voltage dependence of the Ca2+ current; and 4), Ca2+ currents and Ca2+ transients were fully blocked by nifedipine. The loss in voltage-dependent EC coupling promoted by ß2a was inferred by the drastic reduction in maximal Ca2+ fluorescence at large positive potentials (
F/Fmax) in double dysgenic/ß1 KO myotubes overexpressing the pore mutant 1S (E1014K) and ß2a. The data indicate that ß2a, upon interaction with the skeletal pore subunit 1S, overrides critical EC coupling determinants present in 1S. We propose that the 1S/ß pair, and not the 1S-subunit alone, controls the EC coupling signal in skeletal muscle.
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INTRODUCTION
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Excitation-contraction (EC) coupling in striated muscle is a fast process in which a brief depolarization causes an immediate elevation of the cytosolic Ca2+ (Kim and Vergara, 1998
). This process is brought about by a close interaction between the dihydropyridine receptor (DHPR) L-type Ca2+ channel and the ryanodine receptor type 1 (RyR1) in skeletal muscle or type 2 (RyR2) in cardiac muscle. Close proximity between the DHPR and RyR complexes occurs at specialized junctions established between the transverse tubular and sarcoplasmic reticulum (SR) membranes (Franzini-Armstrong and Protasi, 1997). At these junctions, voltage-dependent movements of electrical charges in the skeletal DHPR are coupled to the opening of the RyR1 channel. However, the molecular mechanism that opens the RyR1 channel, including which DHPR subunits directly contribute to this process, is, for the most part, unknown. In contrast, activation of RyR2 channels in cardiac muscle is a Ca2+-dependent process and is strictly proportional to the magnitude of the L-type Ca2+ current (Beuckelmann and Wier, 1988; Bers, 2002). This fundamental difference in EC coupling trigger mechanism leads to significant differences in the voltage-dependence of Ca2+ transients. In skeletal muscle, the Ca2+ transient amplitude versus voltage relationship is sigmoidal whereas in cardiac muscle, it is bell-shaped such that the Ca2+ transient is always proportional to the Ca2+ current. The biophysical and molecular details of SR Ca2+ release mediated by skeletal and cardiac RyR isoforms have been reviewed extensively (McPherson and Campbell, 1993; Coronado et al., 1994; Meissner, 1994; Sutko and Airey, 1996; Fill and Copello, 2002).
In molecular terms, the skeletal DHPR is an 430-kDa heteropentamer composed of 1S, ß1a, 2-
1, and 
1 subunits. The DHPR 1-subunit belongs to the superfamily of voltage-gated channel proteins with four internal repeats. Electrical charges in the S4 segments in each repeat move outward in response to membrane depolarization. This movement is coupled to the opening of the channel (Bezanilla, 2000). ß-subunits are 65-kDa proteins that bind strongly to the cytosolic loop between repeats I and II of 1 via a conserved 30-residue ß interaction domain or BID (Pragnell et al., 1994; De Waard et al., 1994). ß-subunits modulate the kinetics of activation and inactivation of the Ca2+ current (Qin et al., 1996; Wei et al., 2000; Berrou et al., 2001) and strengthen the coupling between S4 charge movements and pore opening (Neely et al., 1993; Olcese et al., 1996; Kamp et al., 1996). The molecular structure and function of the 2-1 and 1 subunits of the skeletal DHPR are far less defined (Gurnett et al., 1996; Ahern et al., 2001c; Arikkath et al., 2003).
Molecular studies of the EC coupling mechanism have relied on myotube models lacking specific DHPR subunits and RyR isoforms. Tanabe et al. (1988) utilized the 1S-null dysgenic myotube to show that the 1S pore subunit was essential for EC coupling signaling. In the dysgenic myotube, a base deletion leads to truncation and degradation of 1S and loss of voltage-activated Ca2+ transients. Transfection of cultured dysgenic myotubes with 1S cDNA leads to de novo synthesis of 1S-subunits (Ahern et al., 2001a,d), concentration of the expressed DHPR in EC coupling junctions (Takekura et al., 1994), and recovery of skeletal type EC coupling (Garcia et al., 1994). This expression system permitted studies in which the missing 1S Ca2+ pore isoform was replaced by 1C, the cardiac/brain pore variant (Tanabe et al., 1990). The Ca2+ fluorescence versus voltage curve expressed by 1C in dysgenic myotubes had a maximum at +30 mV followed by a continuous decrease at more positive potentials. The biphasic nature of this relationship, along with a dependence of Ca2+ transients on external Ca2+, showed that Ca2+-dependent EC coupling could be implemented in skeletal myotubes by a hybrid DHPR composed of a cardiac pore subunit and the nonpore subunits expressed endogenously in the dysgenic myotube. The expression of chimeras of 1S and 1C were later used to demonstrate that a domain within the cytosolic loop linking repeats II and III of 1S (1S L720–Q765) harbored a unique signal essential for skeletal type EC coupling (Nakai et al., 1998a). The studies in dysgenic myotubes have led to the view that the 1S-subunit is the major contributor to the signal transduction and that the 1S II-III loop may be directly responsible for opening RyR1. However, in a strict sense, the observations in dysgenic myotubes cannot exclude the participation of other DHPR subunits in the EC coupling signal since these subunits (2-1, ß, 1) are constitutively expressed in the dysgenic myotube (Arikkath et al., 2003). Hence, if EC coupling domains were to be present in other subunits of the skeletal DHPR, they would have escaped detection in the dysgenic myotube by virtue of being constitutively present. In support of this premise, deletion analysis of the critical 1S L720–Q765 domain suggests that regions other than 1S II-III loop participate in skeletal-type EC coupling (Ahern et al., 2001b).
Gene targeting techniques have substantially increased the number of myotube models that can be used as expression platforms for EC coupling studies. Gregg et al. (1996) described a knockout (KO) of the mouse ß1 gene, encoding DHPR ß isoforms expressed in skeletal muscle (ß1a) and brain (ß1b, ß1c) (Powers et al., 1992). The ß1 KO mutation, like the dysgenic mutation, is perinatally lethal due to the absence of EC coupling in the skeletal musculature. The ß1 KO myotubes fail to contract in response to electrical stimulation despite the presence of normal action potentials, Ca2+ storage capacity, and caffeine-sensitive Ca2+ release. However, ß1 KO cells lack L-type Ca2+ current, and depolarization does not produce Ca2+ transients (Gregg et al., 1996; Strube et al., 1996). Expression of the skeletal muscle ß1a isoform in cultured ß1 KO myotubes resulted in the recovery of the wild-type (WT) L-type Ca2+ current density, the intramembrane charge movement density, and the amplitude and voltage dependence of Ca2+ transients (Beurg et al., 1997). In contrast, expression of the cardiac/brain ß2a variant recovered L-type Ca2+ currents, but the amplitude of depolarization-activated Ca2+ transients was drastically depressed. Because the interaction between the DHPR 1- and ß-subunits is essential for cell surface expression (Chien et al., 1995; Bichet et al., 2000), the loss of EC coupling in the ß1 KO myotube could be exclusively due to the loss of DHPR voltage sensors from the cell surface. Alternatively, domains of the DHPR ß1a-subunit, similar to elements present in the 1S-subunit, might be directly involved in activation of RyR1 channels. To examine the role of DHPR ß1a in skeletal-type EC coupling, we chimerized ß1a and the cardiac/brain ß2a variant and mapped the domain(s) required for functional recovery of skeletal-type EC coupling in cultured ß1 KO myotubes (Beurg et al., 1999a,b; Sheridan et al., 2003a). DHPR 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), a nonconserved amino terminus (D1), and a nonconserved carboxyl terminus (D5) (Perez-Reyez and Schneider, 1994). We tested the participation of D1, D3, and D5 of ß1a in the recovery of Ca2+ conductance, charge movements, and EC coupling in ß1 KO skeletal myotubes (Beurg et al., 1999a; Sheridan et al., 2003a). Deletion of the D5 region of ß1a (ß1a residues 470–524) drastically reduced the amplitude of voltage-evoked Ca2+ transients without affecting the density of DHPR charge movements. Thus, the membrane density of DHPR voltage sensors, critical for voltage-dependent EC coupling, was not compromised by the carboxyl terminal deletion. On the basis of this result, we have implicated the D5 region of ß1a specifically in skeletal-type EC coupling (Sheridan et al., 2003a). Recent recombinant protein binding studies indicate that the D5 domain is essential for strong interaction between DHPR ß1a and a specific cytoplasmic region of RyR1 (Cheng and Coronado, 2003).
Targeted disruption of the three mammalian RyR genes has been described (Takeshima et al., 1994, 1996; Nakai et al., 1996; Bertocchini et al., 1997). Two of these variants, namely RyR1 and RyR3, contribute to EC coupling in skeletal muscle. Absence of RyR1 leads to a full loss of skeletal-type EC coupling. Therefore, the RyR1 KO mutation is also perinatally lethal. Moreover, transient expression of RyR2 or RyR3 in the RyR1-deficient myotube does not lead to a recovery of skeletal EC coupling function (Yamazawa et al., 1996; Nakai et al., 1997; Fessenden et al., 2000). In contrast to RyR1-deficiency, RyR3-deficiency affects dynamic aspects of muscle tension during the embryonic stage and slows Ca2+ transient propagation (Bertocchini et al., 1997; Yang et al., 2001). However, RyR3 is not essential for mouse survival. Such a nonequivalent behavior of RyR1 and RyR3 variants suggests that highly specific interactions between RyR1 and the skeletal DHPR are essential for Ca2+ signaling in skeletal myotubes. This notion has been corroborated by the identification of RyR1 domains involved in skeletal-type EC coupling using chimeras of RyR1 and RyR2 or RyR1 and RyR3 (Yamazawa et al., 1997; Nakai et al., 1998b; Perez et al., 2003).
In this study, we used KO myotubes lacking DHPR ß1, RyR1, and RyR3 to examine the characteristics of the EC coupling induced by the heterologous cardiac/brain ß2a variant in the context of myotubes expressing a WT 1S-subunit. This interest was prompted by the observation that many of the carboxyl terminal deletion mutants of the D5 region of ß1a, when expressed in ß1 KO myotubes, induced Ca2+ transients triggered by the Ca2+ current (Sheridan et al., 2003a). Hence, the EC coupling mechanism induced by the modified skeletal ß1a variants shared many characteristics with the EC coupling process described in the heart. This result begs the question of whether a heterologous DHPR ß variant from a gene predominantly expressed in cardiac tissues might be able to transform the skeletal EC coupling process to one more closely resembling cardiac EC coupling. We show that overexpression of the WT cardiac/brain ß2a variant (Perez-Reyez and Schneider, 1994) produces drastic changes in EC coupling characteristics of cultured skeletal myotubes, altering it from a purely voltage-dependent process to one controlled mostly by the Ca2+ current. The changes are functionally equivalent to those observed in dysgenic myotubes expressing the cardiac/brain 1C variant in the context of the WT DHPR ß1a-subunit. The results are consistent with the view that the DHPR ß-subunit, like 1, is a key structural determinant of EC coupling in muscle cells. Part of this work has been previously published in abstract form (Sheridan et al., 2002, 2003b).
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MATERIALS AND METHODS
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Identification of genotypes
We used polymerase chain reaction (PCR) assays to screen for the WT and mutant alleles of the DHPR 1S, ß1, RyR1, and RyR3 genes in mice with targeted disruptions of these genes (Gregg et al., 1996; Nakai et al., 1996; Bertocchini et al., 1997) or, in the case of muscular dysgenesis mice (mdg), carrying a frame-shifted 1S-null gene (Chaudhari, 1992). Tail samples 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), 1.2 µL Taq polymerase (Qiagen), and 1 µL of the DNA sample (100 µg/mL).
DHPR 1S alleles
PCR primers 5' ttt ccc aca ggc cgt gct gct gct ctt ca 3' and 5' gca gct ttc cac tca gga ggg atc cag tgt 3' were used to amplify a 202-bp fragment of the WT 1S allele. PCR primers 5' gca gct ttc cac tca gga ggg atc cag tgt 3' and 5' ttt ccc aca ggc cgt gct gct gct ctt aga 3' were used to amplify a 203-bp fragment of the mdg allele (1S-null). The following conditions apply for the PCR of both DHPR 1S alleles: 1), 2 min at 94°C; 2), 45 s at 94°C; 3), 45 s at 60°C; 4), 1 min at 72°C; 5), cycle through steps 2–4 for 30 times; and 6), 10 min at 72°C.
DHPR ß1 alleles
PCR primers 5' gag aga cat gac aga ctc agc tcg gag a 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 both DHPR ß1 alleles: 1), 2 min at 94°C; 2), 30 s at 94°C; 3), 45 s at 60°C; 4), 1 min at 72°C; 5), cycle through steps 2–4 for 30 times; and 6), 10 min at 72°C.
RyR1 alleles
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 both RyR1 alleles: 1), 5 min 30 s at 94°C; 2), 1 min at 94°C; 3), 1 min at 57°C; 4), 1 min at 72°C; 5), cycle through steps 2–4 for 35 times; and 6), 10 min at 72°C.
RyR3 alleles
PCR primers 5' cac atc cca atc tcc ttt act cc 3' and 5' gct tat tct gcc cta atg cca c 3' were used to amplify a 316-bp fragment of the WT RyR3 allele. PCR primers 5' ggt cat cct cac ttc gcc tat gtt c 3' and 5' cgt gct act tcc att tgt cac gtc 3' were used to amplify a 920-bp fragment of the KO RyR3 allele. The following conditions apply for the PCR of both RyR3 alleles: 1), 2 min at 94°C (hot start followed by addition of Taq polymerase (Qiagen); 2), 3 min at 94°C; 3), 1 min at 58°C; 4), 2 min at 72°C; 5), cycle through steps 2–4 for 35 times; and 6), 10 min at 72°C.
Primary cultures
Cultures of myotubes were prepared from hind limbs of E18 fetuses, as described previously (Beurg et al., 1997). Muscles dissected from the fetuses were treated 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 myoblast fusion (6 days), the medium was replaced with FBS free medium, and CO2 was decreased to 5%.
Double 1S/ß1-null myotubes
E18 fetuses lacking 1S and ß1a were obtained by screening litters of fetuses from crosses of double heterozygous dysgenic and ß1 KO mice (1S+/mdg, ß1+/- x 1S+/mdg, ß1+/-) as described (Ahern et al., 1999; 2003). Mutant fetuses were visually recognized by the absence of muscle movements and were separately processed for myotube cell culture while a PCR screen was conducted in parallel. Limb myotube cultures from double null E18 fetuses with the genotype 1Smdg/mdg, ß1-/- were utilized in cDNA transfection protocols and are identified in the text as double dysgenic/ß1 KO myotubes.
cDNA transfection
cDNA transfection was performed during the myoblast fusion stage with the polyamine LT1 (Panvera, Madison, WI). Cells were exposed for 2–3 h to a transfection solution containing LT1 and cDNA at a 5:1 µg ratio. In addition to the cDNA of interest, cells were cotransfected with a plasmid encoding the T-cell protein CD8, which is used as a transfection marker. Transfected myotubes expressing CD8 were recognized by surface binding of polystyrene beads coated with a monoclonal antibody specific for an external CD8 epitope (Dynal ASA, Oslo, Norway). The efficiency of cotransfection of the marker and the cDNA of interest was 90%. Whole-cell analysis of Ca2+ currents and Ca2+ transients was performed 3–5 days after transfection.
cDNA constructs
cDNAs for mouse ß1a (residues 1–524; GenBank accession no. NM_031173) and rat ß2a (residues 1–604; GenBank accession no. M80545) were subcloned into the pCR-Blunt vector (Invitrogen, Carlsbad, CA), excised by digestion with AgeI and NotI, and cloned into the pSG5 vector in frame with the first 11 residues of the phage T7 gene 10 protein for antibody tagging. The 1S (E1014K) substitution was introduced in a rabbit 1S template (GenBank accession no. M23919) and was described previously (Ahern et al., 2001b). WT 1S and 1S (E1014K) were fused in-frame to the first 11 amino acids of the phage T7 gene 10 protein in the pSG5 vector.
Whole-cell voltage clamp
Whole-cell recordings were performed as described previously (Strube et al., 1996) with an Axopatch 200B amplifier (Axon Instruments, Foster City, CA). Effective series resistance was compensated up to the point of amplifier oscillation with the Axopatch circuit. All experiments were performed at room temperature. The external solution was (in mM) 130 TEA methanesulfonate, 10 CaCl2, 1 MgCl2, 10-3 TTX, 10 HEPES titrated with TEA(OH) to pH 7.4. The pipette solution consisted of (in mM) 140 Cs aspartate, 5 MgCl2, 0.1 EGTA (when Ca2+ transients were recorded) or 5 EGTA (when only Ca2+ currents were recorded), and 10 MOPS titrated with CsOH to pH 7.2. Patch pipettes had a resistance of 1–1.5 M
when filled with the pipette solution. The limit of Ca2+ current detection was 20 pA/cell or 0.05 pA/pF for the smallest cells having the lowest capacitative noise.
Confocal fluorescence microscopy
Confocal line scan measurements were performed as described previously (Conklin et al., 1999). All experiments were performed at room temperature. Cells were loaded with 5 µM fluo-4 acetoxymethyl (AM) ester (Molecular Probes, Eugene, OR) for 60 min at room temperature. Cells were viewed with an inverted Olympus microscope with a 20x objective (N.A. = 0.4) and a Fluoview confocal attachment (Olympus, Melville, NY). A 488-nm spectrum line for fluo-4 excitation was provided by a 5-mW Argon laser attenuated to 6% with neutral density filters. For voltage-activated Ca2+ transients, line scans consisted of 1000 lines, each of 512 pixels, acquired at a rate of 2.05 milliseconds per line. Line scans were synchronized to start 100 ms before the onset of the depolarization. For Ca2+ transients activated by caffeine or CMC (4-chloro-m-cresol, Sigma, St. Louis, MO), line scans consisted of 1000 lines, each of 512 pixels, acquired at a rate of 30.8 milliseconds per line. Line scans were synchronized to start 800 ms before the onset of the external solution change. The time course of the space-averaged fluorescence intensity change and F/F units were estimated as follows: 1), The pixel intensity in a line scan was transformed into arbitrary units, and the mean intensity of each line was obtained by averaging pixels covering the cell exclusively; 2), The mean resting fluorescence intensity (F) corresponds to the mean intensity of each line averaged before stimulation and was used as a baseline; 3), The change in mean line intensity above baseline (F) was obtained by subtraction of F (baseline) from the mean line intensity; 4), F was divided by the baseline F for each line in a line scan and was plotted as a function of time. The peak-to-peak noise in the baseline fluorescence averaged 0.1 F/F units which was previously estimated to correspond approximately to a 200-nM change in free Ca2+ (Conklin et al., 1999). Since F/F was spatially averaged, noise varied with spatial inhomogeneities in fluo-4 fluorescence and cell size. The smallest fluorescence signals reported in this study were 0.2 F/F in ß2a-overexpressing RyR1/RyR3 KO myotubes and 0.4 F/F in ß2a-overexpressing RyR1 KO myotubes (Table 2). To construct peak Ca2+ fluorescence versus voltage curves, we used the highest F/F line value after the onset of the pulse and up to the termination of the pulse. Image analyses were 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 or 200 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.
Immunostaining
Four to five days after transfection, cells were fixed in 100% methanol and processed for immunostaining as described previously (Gregg et al., 1996). The primary antibody was a mouse monoclonal against the T7 epitope (Novagen, Madison, WI) present in ß1a and ß2a and was used at a dilution of 1:1000. The secondary antibody was a fluorescein conjugated polyclonal goat anti-mouse IgG (Boehringer Mannheim, Indianapolis, IN) and was used at a dilution of 1:1000. Confocal images of 0.3 to 0.4 microns per pixel were obtained in the Olympus Fluoview using a 40x oil-immersion objective (N.A. = 1.3). Images were Kalman-averaged 3 times, and the pixel intensity was displayed as 16 levels of gray in reverse. All images were acquired with minimal laser power (6% of maximum 5 mW) and predetermined PMT settings to avoid pixel saturation and for accuracy in the comparison of images.
Curvefitting
The voltage dependence of the Ca2+ conductance and peak fluorescence, assayed with the 50-ms depolarization, was fitted in all cases according to a Boltzmann distribution (Eq. 1)
 | 1 |
where Amax is Gmax or F/Fmax, V1/2 is the potential at which A = Amax/2, and k is the slope factor. For myotubes overexpressing ß1a, the voltage dependence of peak fluorescence assayed with a 200-ms depolarization was fit with Eq. 1. The ß2a variant produced biphasic bell-shaped fluorescence versus voltage curves with varying degrees of curvature. For myotubes overexpressing ß2a, the voltage dependence of peak intracellular Ca2+ assayed with a 200-ms depolarization was fit with the modified Boltzmann distribution (Eq. 2)
 | 2 |
where (V-Vr) is a factor that accounts for the decrease in Ca2+ current trigger at positive potentials and k' is a scaling factor that varies with the magnitude of F/Fmax. Other parameters are the same as in Eq. 1. Parameters of a fit of averages of many cells (population average) are shown in figures. The statistics of parameters of the fit of individual cells are shown in Table 1. Student's t-tests and analysis of variance (ANOVA) were performed with Analyze-it software (Analyze-it, Leeds, UK).
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RESULTS
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Previous studies showed that ß1 KO myotubes expressing the heterologous DHPR ß2a-subunit had weak Ca2+ transients compared to controls expressing the skeletal muscle-specific ß1a-subunit (Beurg et al., 1999a,b; Sheridan et al., 2002, 2003b). Here we demonstrate that ß2a induces Ca2+ transients triggered by the Ca2+ current, and furthermore, both RyR isoforms described in primary skeletal myotubes are implicated in the functional changes. To separate the contributions of RyR1 and RyR3, we characterized functional expression of ß2a in cultured primary myotubes from RyR1 KO, RyR3 KO, and double RyR1/RyR3 KO mice. Interbreeding mice heterozygous of the RyR1 KO allele and homozygous for the RyR3 KO allele produced RyR1/RyR3 KO mice (Ikemoto et al., 1997; Barone et al., 1998; Conklin et al., 2000). Genotypes were screened by PCR as described in Materials and Methods. Additional controls were performed in ß1 KO myotubes lacking the endogenous ß1a variant. The two DHPR ß variants investigated, namely the heterologous rat ß2a and the homologous mouse ß1a used as a reference, carried a T7 epitope tag fused to the amino terminus for determining relative levels of protein expression in transfected cells. Fig. 1 shows confocal anti-T7 immunofluorescence in four myotube genotypes expressing the homologous and heterologous DHPR ß variants. The cDNA of interest was cotransfected with the CD8 cDNA, which was used to identify viable transfected myotubes in voltage-clamp experiments. Dishes of transfected cells were incubated with anti-CD8 antibody beads, fixed, and processed for immunostaining with anti-T7 antibody. Protein expression was determined 4–5 days after transfection. Voltage-clamp experiments have determined that this transfection time is adequate for full functional recovery of EC coupling properties in ß1 KO myotubes (Beurg et al., 1997). At this stage of protein expression, we found abundant ß-protein throughout the myotube with the exclusion of the cell nuclei. Based on fluorescence intensity, we estimated that all myotube genotypes overexpressed exogenous ß2a and ß1a proteins at comparable levels.
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FIGURE 1 ß-protein expression in ß1 KO and RyR KO myotubes. Confocal immunofluorescence of myotubes expressing T7-tagged DHPR ß1a- or ß2a-subunits. Cells with the indicated genotype were transfected with CD8 cDNA plus the ß1a or ß2a cDNA. ß1a corresponds to the full-length mouse cDNA; ß2a corresponds to the full-length rat cDNA. Cells were incubated with CD8 antibody beads, fixed, and stained with anti-T7 primary/fluorescein-conjugated secondary antibodies. Pixel intensity was converted to a 16-level inverted gray scale with high-intensity pixels in black color. Images are 84 x 56 microns and on-focus beads are 4.5 microns in diameter. The asterisk indicates a nontransfected myotube in the same focal plane as the transfected cell.
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Transcripts for RyR3 have been detected in primary cell cultures of WT and RyR1 KO limb myotubes (Takeshima et al., 1995). Furthermore, functional RyR3 channels account for caffeine-induced Ca2+ release and Ca2+ sparks detected in embryonic RyR1 KO myotubes (Takeshima et al., 1995; Conklin et al., 2000). In contrast, RyR3 has been reported to be absent in a myogenic cell line derived from RyR1 KO embryonic limb myotubes (Fessenden et al., 2000). To determine whether one or two RyR variants were present in our primary cell cultures, we conducted Ca2+ release studies using caffeine and the high-affinity RyR1 agonist CMC (Herrmann-Frank et al., 1996). In these experiments, cells were loaded with fluo-4 AM, and SR Ca2+ release was induced by fast perfusion of external solution supplemented with caffeine (10 mM) or CMC (0.5 mM). The perfusion solution was passed through a large-bore pipette placed near the cell with the pipette connected to a pressurized manifold (ALA Scientific, Westbury, NY) and synchronized to confocal line scan image acquisition. Fig. 2 shows Ca2+ transients evoked by fast perfusion of caffeine or CMC in WT, RyR3 KO, RyR1 KO, and RyR1/RyR3 KO myotubes. As indicated by the line diagram above each trace of fluorescence, the external solution change lasted 4 s and was initiated 800 ms after the start of the line scan. After the pulse of agonist, line scan acquisition was continued for an additional 26 s. The time course of the space-averaged confocal fluorescence intensity was computed from the line scan image and is shown in F/F units. A rapid increase in myotube fluorescence was observed in response to caffeine in WT and RyR3 KO myotubes. Comparatively weaker and slower responses to caffeine were present in 60% of the tested RyR1 KO myotubes (26 of 41 cells) with the rest entirely unresponsive to caffeine. Caffeine had no effect in double RyR1/RyR3 KO myotubes (14 of 14 cells). The presence of caffeine-sensitive Ca2+ pools in RyR1 KO and RyR3 KO, but not in RyR1/RyR3 KO myotubes, confirmed the presence of these two RyR isoforms in our cell cultures and was consistent with previous studies (Takeshima et al., 1995; Conklin et al., 2000). However, since the response to caffeine in RyR1 KO myotubes was heterogeneous, functional RyR3 channels may not be present in all RyR1-deficient cells. Alternatively, RyR3 channels may not be responsive to caffeine in all cells for some unknown reason. Fig. 2 also shows that large and fast responses to CMC were observed in WT and RyR3 KO myotubes, and much weaker and slower responses were present in RyR1 KO and double RyR1/RyR3 KO myotubes. The results in RyR3 KO compared to RyR1 KO myotubes agreed with previous determinations showing that CMC is predominantly a RyR1 agonist (Fessenden et al., 2000). Yet, the presence of a small but consistent response to CMC in double RyR1 KO/ RyR3 KO myotubes (15 of 15 cells) implicates targets other than RyRs, at least in myotubes in which both RyR isoforms are absent. Histograms of the mean maximal fluorescence induced by caffeine and CMC and the time to maximal fluorescence after agonist exposure are shown at the bottom of Fig. 2. Asterisks indicate ANOVA significance p < 0.05 relative to WT. The absence of a response to caffeine in double KO myotubes provided a null background against which the responses in RyR1 KO and RyR3 KO myotubes could be compared. We observed a higher peak and faster time-to-peak responses to caffeine in RyR3 KO than in RyR1 KO myotubes, relative to the null background. This suggested that RyR3 KO cells had a higher sensitivity to caffeine than RyR1 KO cells. Alternatively, RyR3 KO cells expressed a higher density of RyR1 receptors than the density of RyR3 receptors expressed in RyR1 KO cells. Since 10 mM caffeine was reported to saturate responses in myotubes expressing either RyR1 or RyR3 (Fessenden et al., 2000), it is likely that the differential sensitivity to caffeine reflects differences in the functional density of RyRs with RyR1 present at a much higher density than RyR3, respectively, in RyR3 KO and RyR1 KO myotubes. It is also possible that RyR1 expression may be up-regulated in RyR3 KO myotubes relative to WT since CMC responses were significantly higher in the former than the latter. In summary, the Ca2+ release responses to caffeine and CMC show that functional RyR1 and RyR3 are present, respectively, in RyR3 KO and most RyR1 KO cultured myotubes.
Fig. 3 shows Ca2+ conductance versus voltage relationships in myotubes transfected with each of the two DHPR ß isoforms (ß1a and ß2a in top and bottom rows, respectively) in each of four genotypes (ß1 KO, RyR3 KO, RyR1 KO, and RyR1/RyR3 KO in columns from left to right). ß2a-transfected myotubes were compared to ß1a-transfected myotubes even though both RyR KO genotypes are likely to express a native DHPR that includes ß1a. We felt that in the case of the RyR KO genotypes, myotubes with exogenous ß1a overexpression provided a better control than nontransfected myotubes since overexpression eliminated potential phenotypic changes introduced by an unknown up- or down-regulation of endogenous ß1a. However, this concern was largely unfounded. The insets show whole-cell Ca2+ currents elicited by 500-ms depolarizations to -10 and +30 mV from a holding potential of -40 mV. The complete pulse protocol utilized for fitting conductance versus voltage curves consisted of 500-ms step depolarizations from -35 mV to +60 mV every 5 mV. The gray lines in each plot indicate the mean conductance of nontransfected myotubes. We found that nontransfected ß1 KO myotubes had marginally detectable Ca2+ currents in agreement with previous studies (Ahern et al., 2003). In ß1a-expressing ß1 KO myotubes, we consistently detected a high-density Ca2+ current with a maximal conductance, Gmax, and other Boltzmann parameters, similar to those of WT myotubes (Table 1). Thus, as shown previously (Beurg et al., 1997), ß1a overexpression in the ß1 KO myotube recovered the normal Ca2+ current phenotype. Furthermore, the Ca2+ current density of WT myotubes was similar to that reported previously by others and us (Beurg et al., 1997; Strube et al., 1998; Avila and Dirksen, 2000). Averages of Boltzmann parameters fitted to the conductance versus voltage curve of each cell and the statistical significance of the data are shown in Table 1. Overexpression of ß1a in the RyR3 KO myotube did not increase Ca2+ current density beyond that present in nontransfected RyR3 KO myotubes, which was entirely normal (Gmax = 182 ± 15 pS/pF, n = 4 cells versus 162 ± 15 pS/pF, n = 9 cells for nontransfected and ß1a-transfected, respectively). In RyR1 KO and RyR1/RyR3 KO myotubes, the endogenous Ca2+ current density was severely depressed, and ß1a overexpression failed to increase it beyond levels present in nontransfected cells. The Gmax of nontransfected RyR1 KO and RyR1/RyR3 KO myotubes were 27 ± 5 pS/pF (14 cells) and 24 ± 10 pS/pF (4 cells), respectively. These values were statistically similar to the density of ß1a-transfected RyR1 KO and RyR1 KO/RyR3 KO myotubes shown in Table 1. The inability of exogenous ß1a overexpression to increase the Ca2+ current density in RyR1-deficient myotubes was not surprising in light of the known fact that RyR1 is a strong enhancer of the native DHPR Ca2+ current (Nakai et al., 1996; Avila et al., 2001; Ahern et al., 2003). Thus, absence of RyR1, and not the endogenous level of the DHPR ß1a-subunit, may be the limiting factor for Ca2+ current expression in the RyR1-deficient genotypes. In ß1 KO myotubes, ß2a expressed a high-density Ca2+ current with a Gmax similar to that recovered by ß1a (Table 1). Moreover, the half-activation potential and voltage dependence of the Ca2+ current expressed by the ß1a and ß2a were also similar. Also, previous studies had shown that Ca2+ currents recovered by ß1a and ß2a had similar mean-variance noise characteristics and activation kinetics (Beurg et al., 1999b). Hence it is highly likely that ß2a became integrated into a functional skeletal DHPR with the pore subunit 1S providing the pathway for the Ca2+ current. The fate of the two other subunits of the DHPR in this hybrid complex is not known. However, their presence in the complex is suggested by the complete normalcy of the Boltzmann parameters of the Ca2+ conductance recovered by ß2a (Table 1). ß2a overexpression in RyR1 KO myotubes was recently shown to restore a slow Ca2+ current with an entirely WT density (Ahern et al., 2003). The bottom row of Fig. 3 confirmed this observation and further showed that ß2a recovered high-density Ca2+ currents in all genotypes analyzed, including double RyR1/RyR3 KO myotubes. In the RyR1 KO and RyR1 KO/RyR3 KO myotubes, ß2a recovered Ca2+ currents with a density 5 fold larger than that present in nontransfected cells and similar to that of WT myotubes (Table 1). Thus, in contrast to the behavior of ß1a, Ca2+ currents recovered by ß2a readily bypassed the inhibition imposed by the absence of RyR1. Furthermore, the results in double KO myotubes showed that neither RyR1 nor RyR3 were required for DHPR Ca2+ current expression when the heterologous ß2a-subunit was present in the DHPR complex. Given this drastic change in Ca2+ current expression pattern, we further tested if ß2a was capable of recruiting a pore subunit other than 1S which could potentially be present in the cultured myotube and that could account for the Ca2+ currents recovered by ß2a. Controls indicated that neither ß1a nor ß2a recovered Ca2+ currents when overexpressed in 1S-null dysgenic myotubes in similar cell culture conditions (not shown). Thus, non-1S pore subunits, if present in skeletal myotubes, may not be able to combine with ß1a or ß2a to generate functional Ca2+ channels. This result strongly suggested that Ca2+ currents recovered by ß2a in RyR1 KO and RyR1/RyR3 KO myotubes originated from hybrid DHPR complexes that included 1S and the exogenous ß2a-subunit. A mechanism explaining how Ca2+ currents recovered by ß2a bypass the requirement for RyR1 has been proposed elsewhere (Ahern et al., 2003).
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FIGURE 3 Ca2+ conductance expressed by the heterologous DHPR ß2a-subunit in ß1 KO and RyR KO myotubes. Myotubes with the indicated genotype (ß1 KO, RyR3 KO, RyR1 KO, and double RyR1/RyR3 KO) were transfected with ß1a (top row) or ß2a (bottom row). Insets show representative Ca2+ currents at -10 mV and +30 mV for a depolarization of 500 ms from a holding potential of -40 mV. Lines correspond to a Boltzmann fit of the population mean Ca2+ conductance indicated in Table 1. All conductance versus voltage curves were fit with Eq. 1. Parameters of the fitted lines, with Gmax in pS/pF, V1/2 in mV, and k in mV, were as follows: for ß1 KO myotubes, were 194, 14.9, and 7 for ß1a overexpression and 183, 11.6, and 5.2 for ß2a overexpression; for RyR3 KO myotubes, were 162, 15.5, 6.1 for ß1a and 156, 10.5, 5 for ß2a; for RyR1 KO myotubes were 25, 17.9, 6.6 for ß1a and 184, 13.2, 6.4 for ß2a; for RyR1/RyR3 KO myotubes were 35, 25.8, 5.8 for ß1a and 157, 14.6, 8 for ß2a.
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In ß1 KO myotubes overexpressing ß1a, the Ca2+ current has no bearing on the magnitude of the Ca2+ transient (Beurg et al., 1997). However, recent studies showed that long depolarizations significantly increased the total Ca2+ entering the cell, and that the excess of Ca2+ entry triggered SR Ca2+ release when ß1 KO myotubes overexpressed truncated variants of ß1a (Sheridan et al., 2003a). For this reason, possible changes in EC coupling produced by ß2a overexpression were tested by voltage steps with a duration of 50 ms and 200 ms in the range of -30 mV to +90 mV, which covered both the inward and outward phases of the myotube Ca2+ current. Fig. 4 shows confocal Ca2+ transients in response to 200-ms depolarizations to +30 and +90 mV from a holding potential of -40 mV in the same myotube. Ca2+ currents were acquired concurrently, and they were similar in density to those described in Fig. 3. The entire pulse protocol consisted of 50-ms and 200-ms depolarizations, and a comparison of Ca2+ transients stimulated by the two protocols is described below. Because the Ca2+ current becomes progressively smaller at potentials >30 mV, as the imposed voltage approaches the Ca2+ current reversal potential and the Ca2+ equilibrium potential, a contribution of the Ca2+ current as trigger of the Ca2+ transient can be readily deduced by comparing Ca2+ transients at +30 mV and +90 mV. Fig. 4 shows that in ß1 KO cells overexpressing ß1a, Ca2+ transients had a nearly identical shape and magnitude at +30 and +90 mV. This result was consistent with the previous data (Beurg et al., 1999a,b; Sheridan et al., 2003a) and the well-known voltage dependence of skeletal-type EC coupling, which is unrelated to the fate of the Ca2+ current (Rios and Pizarro, 1991). Similarly, Ca2+ transients in ß1a-expressing RyR3 KO myotubes had the same maximal intensity at +30 and +90 mV, and this result agreed with previous determinations showing that RyR3 KO myotubes display skeletal-type EC coupling (Dietze et al., 1998). In myotubes lacking RyR1 (RyR1 KO and RyR1/RyR3 KO), ß1a overexpression failed to recover Ca2+ transients, which is not surprising since RyR3 is incapable of supporting skeletal-type EC coupling (Fessenden et al., 2000). Additionally, the Ca2+ current density in the RyR1-deficient genotypes is noticeably small. Hence, Ca2+-dependent Ca2+ release or a direct contribution of the Ca2+ current to the cell fluorescence would be difficult to detect in these cells. In contrast, Ca2+ transients of various magnitudes were detected in all myotube genotypes overexpressing ß2a and were two- to fivefold smaller at +90 than at +30 mV (Fig. 4, bottom row). Furthermore, in RyR1 KO and RyR1/RyR3 KO myotubes, ß2a-mediated Ca2+ transients were seen at +30 but not at +90 mV, suggesting that a significant proportion of the fluorescence signal in these myotubes could arise from the Ca2+ current itself. ß2a-mediated Ca2+ transients in ß1 KO myotubes were smaller in magnitude than those in ß1a-expressing ß1 KO, in agreement with previous determinations (Beurg et al., 1999b). In addition, the selected ß2a-expressing RyR3 KO myotube shows Ca2+ transients larger than that of the ß2a-expressing ß1 KO myotube. However, on average, this difference was not significant (Table 2). The results in ß1 KO and RyR3 KO myotubes suggested that ß2a expression changed the EC coupling signal from one that is controlled by voltage to one in which the Ca2+ current served as a trigger. We were especially surprised by the changes in the voltage dependence of Ca2+ transients in RyR3 KO myotubes since nontransfected RyR3 KO myotubes (Dietze et al., 1998) or ß1a-overexpressing RyR3 KO myotubes (Fig. 4, top row) showed bona fide skeletal-type EC coupling. This result seemed to indicate that ß2a behaved as a negative dominant subunit capable of disrupting normal DHPR function in the RyR3 KO myotube.
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FIGURE 4 Ca2+ transients expressed by the heterologous DHPR ß2a-subunit in ß1 KO and RyR KO myotubes. Myotubes with the indicated genotype (ß1 KO, RyR3 KO, RyR1 KO, and double RyR1/RyR3 KO) were transfected with ß1a (top row) or ß2a (bottom row). Ca2+ transients were elicited by step depolarizations to +30 and +90 mV. Myotubes were held at a resting potential of -40 mV and depolarized for 200 ms. The time course of the cell fluorescence was obtained by integration of the line scan image as described in Materials and Methods. Note differences in F/F scales.
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To determine how much of the fluorescence signal in ß2a-expressing myotubes was due to the Ca2+ current and how much was due to SR Ca2+ release, we examined fluorescence versus voltage and Ca2+ current versus voltage relationships acquired concurrently. Fluorescence signals in RyR1/RyR3 KO myotubes should exclusively reflect the contribution of the Ca2+ current, whereas those in RyR1 KO should reflect the contribution of the Ca2+ current and Ca2+ entry dependent Ca2+ release mediated by RyR3. Consistent with this idea, Fig. 5 A shows that ß2a-mediated Ca2+ transients in RyR1 KO (closed circles) and RyR1 KO/RyR3 KO (closed squares) myotubes had a bell-shaped dependence with a maximum at +30 mV, which coincided with the maximal Ca2+ current shown in Fig. 5 B. Additionally, the peak F/F at +30 mV was larger in RyR1 KO than in RyR1 KO/RyR3 KO myotubes, whereas the Ca2+ current density was similar in both cases. The baseline for these curves was provided by the behavior of nontransfected RyR1 KO myotubes (open circles) and RyR1/RyR3 KO myotubes (open squares), which lacked Ca2+ transients entirely and expressed a minimal Ca2+ current. The peak F/F of nontransfected myotubes was undistinguishable from the noise of the measurement, which was 0.1 F/F (see Materials and Methods). We surmise that the excess in cell fluorescence in ß2a-expressing RyR1 KO myotubes, above that present in ß2a-expressing RyR1 KO/RyR3 KO myotubes, should reflect SR Ca2+ release due to Ca2+-dependent activation of RyR3. To confirm this idea, ß2a-expressing RyR1 KO myotubes were treated with 5 µM ryanodine to block any RyR-dependent SR Ca2+ release. Fig. 5 C shows representative Ca2+ transients and Ca2+ currents at +30 mV in a ß2a-expressing RyR1 KO myotube (black traces) and in a separate ß2a-expressing RyR1 KO myotube treated with ryanodine (gray traces). To ensure a complete elimination of the fluorescence change due to EC coupling, myotubes were exposed to ryanodine for 30 min. For this reason, controls and ryanodine-treated cells were different. Myotubes were selected so that the Ca2+ current densities were approximately the same. Ryanodine treatment significantly reduced the fluorescence signal in ß2a-expressing RyR1 KO myotubes and suggested that a significant component of the fluorescence signal in ß2a-expressing RyR1 KO myotubes was due to RyR-dependent SR Ca2+ release presumably mediated by RyR3. Since Ca2+ current densities in all ß2a-expressing genotypes were roughly similar (Table 1), we estimated the relative magnitude of the ß2a-mediated Ca2+ release in all genotypes by normalizing the peak F/F relative to the magnitude of the Ca2+ current at the same potential of +30 mV (Fig. 5 D). The normalized signal in RyR1/RyR3 KO myotubes represents the direct contribution of the Ca2+ current to the cell fluorescence, and progressively larger signals due to SR Ca2+ release were present in RyR1 KO, RyR3 KO, and ß1 KO myotubes. The largest Ca2+ release signals were observed in the RyR1-expressing myotubes (ß1 KO and RyR3 KO) consistent with the idea that RyR1 is expressed at a much higher density than RyR3 described above. Voltage-dependent components in ß2a-transfected RyR3 KO and ß1 KO myotubes also contributed to the fluorescence signal in these cells and are discussed separately (see Figs. 6 and 9). The cell fluorescence to Ca2+ current ratio at +30 mV was not statistically different for ß2a-transfected RyR3 KO and ß1 KO myotubes. However, this ratio was significantly smaller for ß2a-transfected RyR1 KO and RyR1/RyR3 KO myotubes (t-test significance p < 0.05) and is indicated by asterisks. The main conclusions from the comparison in Fig. 5 D are that the Ca2+ current has a modest impact on the measured myotube fluorescence and that both RyR isoforms (RyR1 and RyR3), when present, can be activated by the Ca2+ current recovered by ß2a overexpression.
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FIGURE 6 Sigmoidal and bell-shaped voltage dependence of Ca2+ transients in myotubes expressing the heterologous DHPR ß2a-subunit. Ca2+ transients in F/F units were measured in myotubes with the indicated phenotype (ß1 KO, RyR3 KO, RyR1 KO and double RyR1/RyR3 KO) expressing ß1a (top row) or ß2a (bottom row). The depolarization duration was 50 ms (closed symbols) or 200 ms (open symbols) from a holding potential of -40 mV. The same myotube was subjected to the 50-ms and 200-ms depolarization protocols. Ca2+ transients were measured at the end of each depolarization from the time course of the integrated confocal line scan image fluorescence. The lines correspond to a Boltzmann fit of the population mean F/F indicated in Table 2. All fluorescence vs. voltage curves were fit with Eq. 1 except those obtained with the 200 ms depolarization in myotubes overexpressing ß2a, which were fit with Eq. 2. Parameters of the fitted lines (F/Fmax in F/F units, V1/2 in mV, and k in mV) for ß1 KO overexpressing ß1a are: 2.8, -6.1, 7.8 for 50 ms and 3.2, -10.2, 5.7 for 200 ms. Parameters for ß1 KO overexpressing ß2a are: 0.8, -2.5, 5.8 for 50 ms and 1.8, 9, 9.7 for 200 ms. Parameters for RyR3 KO overexpressing ß1a are: 2.2, -7.2, 8.2 for 50 ms and 3.2, -9.9, 7.3 for 200 ms. Parameters for RyR3 KO overexpressing ß2a are: 0.6, 8.5, 11.3 for 50 ms and 1.5, 3.6, 10 for 200 ms. Parameters for RyR1 KO overexpressing ß2a are: 0.4, 9.8, 8.9 for 200 ms. Parameters for RyR1/RyR3 KO overexpressing ß2a are: 0.2, 7.9, 10 for 200 ms.
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FIGURE 9 Ca2+ transients and Ca2+ currents expressed by the heterologous DHPR ß2a-subunit in 1S/ß1-null myotubes. Ca2+ transients and Ca2+ currents in the same myotube at +30 mV from a holding potential of -40 mV in response to a 200-ms depolarization. Panel A shows a ß1 KO myotube transfected with ß1a (black traces) and a dysgenic myotube (labeled mdg) transfected with 1S (E1014K) (gray traces). Panel B shows average fluorescence versus voltage and current versus voltage curves for ß1 KO myotubes expressing ß1a (black traces) and dysgenic myotubes expressing 1S (E1014K) (gray traces). Fluorescence and whole cell current data are shown for 10 and 5 cells in black and gray symbols, respectively. Panel C shows a ß1 KO myotube transfected with ß2a (black traces) and a double dysgenic /ß1 KO myotube cotransfected with ß2a and 1S (E1014K) (gray traces). Panel D shows average fluorescence versus voltage and current versus voltage curves for ß1 KO myotubes expressing ß2a (black traces) and for double dysgenic/ß1 KO myotubes coexpressing ß2a + 1S (E1014K) (gray traces). Fluorescence and whole cell current data are shown for 10 and 8 cells in black and gray symbols, respectively. Parameters of the Boltzmann fit of fluorescence were F/Fmax in F/F units, V1/2 in mV, and k in mV) were 4 ± 0.6, 1 ± 6, 8 ± 1.1 (1S (E1014K), 5 cells) and 0.3 ± 0.1, 24 ± 5, 9.3 ± 1.3 (ß2a + 1S (E1014K), 8 cells). Parameters for ß1a and ß2a expressing cells in ß1 KO myotubes are shown in Table 2.
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Ca2+ release triggered by the Ca2+ current follows the voltage dependence of the Ca2+ current. Hence, this EC coupling mechanism can be distinguished from purely voltage-dependent EC coupling by the shape of the Ca2+ fluorescence versus voltage relationship. Additionally, Ca2+-dependent EC coupling should be a sensitive function of the duration of the depolarizing stimulus as longer depolarizations increase the amount of trigger Ca2+ entering the cell. Previous studies showed that depolarizations for 50 ms were adequate for activation of DHPR charge movements (Ahern et al., 2003). However, activation of the Ca2+ current to near steady state required depolarizations of 200 ms (Sheridan et al., 2003a). For these reasons, we investigated the shape of fluorescence versus voltage curves for depolarizations of 50 and 200 ms in the same cell. Fluorescence versus voltage relationships are shown in Fig. 6 for ß1a and ß2a overexpression in the four myotube genotypes under investigation. The F/F value reached at the end of the depolarization was plotted as a function of voltage for depolarizations of 50 ms (closed symbols) and 200 ms (empty symbols). The gray traces indicate the mean fluorescence of nontransfected myotubes. The duration of the pulse did not affect the sigmoidal voltage dependence of Ca2+ transients expressed by ß1a in ß1 KO myotubes, which is in agreement with recent observation from our laboratory (Sheridan et al., 2003a) and studies in adult skeletal muscle (Melzer et al., 1986). Interestingly, RyR3 KO myotubes overexpressing ß1a had sigmoidal fluorescence versus voltage curves with a higher maximal fluorescence (F/Fmax) for the 200 ms than for the 50 ms stimulation. This difference was also observed in nontransfected myotubes (F/Fmax = 1.8 ± 0.1 for 50 ms and 2.7 ± 0.1 for 200 ms for four cells) and is shown by the two gray traces in the top and bottom graphs corresponding to RyR3 KO myotubes. However, the differences in F/Fmax for 50-ms and 200-ms depolarizations were not significant either in ß1a-transfected (see Table 2) or nontransfected RyR3 KO myotubes. A contribution of RyR3 to the overall rate of propagation of Ca2+ release has been previously reported in cultured RyR3 KO myotubes and could underlie the small difference in F/Fmax observed here for the short and long depolarizations (Yang et al., 2001). In RyR1 KO and RyR1/RyR3 KO myotubes, ß1a overexpression failed to recover Ca2+ transients entirely, in agreement with the critical role of RyR1 in voltage-dependent coupling and the absence of ß1a stimulated Ca2+ current expression in these genotypes. ß2a overexpression changed the shape of the fluorescence versus voltage curve in the RyR1-expressing genotypes (ß1 KO and RyR3 KO) and introduced Ca2+ transients in the RyR1-deficient genotypes (RyR1KO and RyR1/RyR3 KO) that were not present in nontransfected counterparts. In ß1 KO and RyR3 KO myotubes, ß2a-overexpression reduced the F/Fmax produced by the 50-ms stimulation relative to that recovered by ß1a, and furthermore, the fluorescence versus voltage curves generated with the 200-ms pulse were clearly bell-shaped. Bell-shaped curves were also observed for the 200-ms depolarization in RyR1 KO and RyR1/RyR3 KO myotubes but not for the 50-ms depolarization. In all cases, the 200-ms depolarization produced an increase in fluorescence in the range of -30 mV to +20 mV, a maximum at +30 mV, and a decline in fluorescence in the range of +40 mV to +90 mV. To quantify these observations, we fitted the curves obtained with the 200-ms depolarization for ß2a-expressing myotubes with a modified Boltzmann equation that takes into account the reversal potential of the Ca2+ current (Eq. 2), whereas the rest of the curves in Fig. 6 were fit with a conventional Boltzmann equation (Eq. 1). The degree of curvature of the fluorescence versus voltage plots at positive potentials was estimated by computing the ratio of fluorescence at the experimental maximum (exp max) for a given fluorescence versus voltage curve (typically +30mV) and the fluorescence at +90 mV 
. The Boltzmann parameters of the fit of each cell and the fluorescence ratios at 50 ms and 200 ms are shown in Table 2 along with the statistical significance and the number of cells. In summary, a dependence of the Ca2+ transient amplitude on pulse duration was observed for ß2a-overexpressing myotubes. This was reflected in the bell-shaped fluorescence versus voltage relationships, consistent with the presence of an EC coupling component triggered by the Ca2+ current in all myotube genotypes overexpressing ß2a. Ca2+ transients were the smallest in RyR1/RyR3 KO myotubes, and, as mentioned previously, they reflected the fluorescence contributed by the Ca2+ current itself.
If the Ca2+ current triggered SR Ca2+ release in myotubes expressing ß2a, the rate of rise of the fluorescence signal and the rate of Ca2+ entering the cell should have a similar time course. Fig. 7 shows Ca2+ transients and Ca2+ currents produced by a 200-ms depolarization to +30 mV from a holding potential of -40 mV in the four myotube genotypes. To correlate the time course of the Ca2+ transient and the Ca2+ current, we compared the cumulative sum of the Ca2+ entry charge during the depolarization (i.e., the running integral of the Ca2+ current) with the fluorescence signal. We have previously argued that this is a fair comparison because Ca2+ fluorescence was measured in an internal solution containing a low EGTA concentration (0.1 mM), and, thus, the cell fluorescence tracked Ca2+ accumulation rather than the rate of SR Ca2+ release (Sheridan et al., 2003a,b). Accordingly, the confocal fluorescence every 2.05 ms (gray dots in F/F units), and the running integral of the Ca2+ current (smooth curve in black in units of pA x ms) were superimposed. For the RyR1-expressing myotubes (ß1 KO and RyR3 KO) with ß1a-overexpression, the fluorescence signal increased earlier and faster than the Ca2+ charge. Furthermore, the fluorescence signal saturated before termination of the200-ms pulse, whereas the Ca2+ charge continued to increase for as long as the pulse was at +30 mV. In contrast, Ca2+ transients obtained by ß2a-overexpression were considerably slower than those obtained by ß1a-overexpression. Furthermore, there was a much closer agreement in the kinetics of the two signals in myotubes overexpressing ß2a, which was particularly compelling in the ß2a-expressing RyR1 KO myotube. In RyR1/RyR3 KO myotubes, the fluorescence signal was too noisy to permit a comparison, and, for that reason, it was not expanded. In summary, ß2a overexpression eliminated the fast EC coupling typical of skeletal myotubes. Furthermore, the slow EC coupling observed in myotubes expressing ß2a appears to have its kinetic basis on the slow time course of the skeletal DHPR Ca2+ current.
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FIGURE 7 Correlation between the rate of cytosolic Ca2+ increase and rate of Ca2+ accumulation in myotubes overexpressing the heterologous DHPR ß2a-subunit. Each panel shows Ca2+ currents and Ca2+ transients in myotubes of the indicated genotype (ß1 KO, RyR3 KO, RyR1 KO, and double RyR1/RyR3 KO) overexpressing ß1a (top row) or ß2a (bottom row) in response to a 200-ms depolarization from a holding potential of –40 mV to +30 mV in external solution with 10 mM Ca2+. The digitized trace (gray) shows fluo-4 fluorescence in F/F units during the confocal line scan. Each dot corresponds to the mean fluorescence of a single 512-pixel line acquired at a rate of 2.05 ms per line. The black trace shows the cumulative integral of the Ca2+ current (running integral) in fC (pA x ms) superimposed on the fluorescence trace. Actual Ca2+ currents are shown below the trace of fluorescence. Note changes in F/F | |
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ABSTRACT
INTRODUCTION
