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Multiple Loops of the Dihydropyridine Receptor Pore Subunit Are Required for Full-Scale Excitation-Contraction Coupling in Skeletal Muscle

Department of Physiology, University of Wisconsin School of Medicine, Madison, Wisconsin

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.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Understanding which cytosolic domains of the dihydropyridine receptor participate in excitation-contraction (EC) coupling is critical to validate current structural models. Here we quantified the contribution to skeletal-type EC coupling of the 1S (Ca_V1.1) II-III loop when alone or in combination with the rest of the cytosolic domains of 1S. Chimeras consisting of 1C (Ca_V1.2) with 1S substitutions at each of the interrepeat loops (I-II, II-III, and III-IV loops) and N- and C-terminal domains were evaluated in dysgenic (1S-null) myotubes for phenotypic expression of skeletal-type EC coupling. Myotubes were voltage-clamped, and Ca²⁺ transients were measured by confocal line-scan imaging of fluo-4 fluorescence. In agreement with previous results, the 1C/1S II-III loop chimera, but none of the other single-loop chimeras, recovered a sigmoidal fluorescence-voltage curve indicative of skeletal-type EC coupling. To quantify Ca²⁺ transients in the absence of inward Ca²⁺ current, but without changing the external solution, a mutation, E736K, was introduced into the P-loop of repeat II of 1C. The Ca²⁺ transients expressed by the 1C(E736K)/1S II-III loop chimera were 70% smaller than those expressed by the Ca²⁺-conducting 1C/1S II-III variant. The low skeletal-type EC coupling expressed by the 1C/1S II-III loop chimera was confirmed in the Ca²⁺-conducting 1C/1S II-III loop variant using Cd²⁺ (10^–4 M) as the Ca²⁺ current blocker. In contrast to the behavior of the II-III loop chimera, Ca²⁺ transients expressed by an 1C/1S chimera carrying all tested skeletal 1S domains (all 1S interrepeat loops, N- and C-terminus) were similar in shape and amplitude to wild-type 1S, and did not change in the presence of the E736K mutation or in the presence of 10^–4 M Cd²⁺. Controls indicated that similar dihydropyridine receptor charge movements were expressed by the non-Ca²⁺ permeant 1S(E1014K) variant, the 1C(E736K)/1S II-III loop chimera, and the 1C(E736K)/1S chimera carrying all tested 1S domains. The data indicate that the functional recovery produced by the 1S II-III loop is incomplete and that multiple cytosolic domains of 1S are necessary for a quantitative recovery of the EC-coupling phenotype of skeletal myotubes. Thus, despite the importance of the II-III loop there may be other critical determinants in 1S that influence the efficiency of EC coupling.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Ca²⁺ signals of skeletal muscle cells are controlled by the voltage-gated L-type Ca²⁺ channel formed by the dihydropyridine receptor (DHPR), and by the sarcoplasmic reticulum (SR) Ca²⁺ release channel formed by the ryanodine receptor type 1 (RyR1). The well-established paradigm is that in response to depolarization, the DHPR produces a signal that briefly opens the RyR1 channel, leading to the release of SR-stored Ca²⁺. Signal transmission takes place at specialized junctions between transverse tubules and SR membranes. At these junctions, DHPRs in tetrad arrangement juxtapose a single tetrameric RyR1 channel (1–3). The narrow physical gap between the transverse tubule and the SR (12 nm) (2), the large protrusion of foot structure of the RyR1 channel into the gap (7 nm) (4), and the overall molecular dimensions of the DHPR and RyR1 complexes (5,6), all suggest that the DHPR and RyR1 channels must be in physical contact. Strong evidence for the formation of a DHPR-RyR1 complex in myotubes is provided by a recent freeze-fracture analysis of the molecular determinants that specify the arrangement of DHPRs in arrays of tetrads opposite to RyR1 (7).

The 1S subunit (Ca_V1.1) of the skeletal DHPR has the familiar topology of a four-repeat voltage-gated channel with five cytosolic domains adjoining the four repeats (N-terminus, I-II loop, II-III loop, III-IV loop, and C-terminus) (8–10). Reports made almost 15 years ago (11,12) and refined later (13–17), have suggested that the cytosolic loop linking repeats II and III, consisting of 132 residues, brings about the conformational change that opens the RyR1 channel under the influence of membrane depolarization. The II-III loop is commonly viewed as the cell's version of the mechanical plunger proposed by Chandler et al. (18), in which an element of the transverse tubule linked to the excitation-contraction (EC) -coupling voltage sensor exerts torque on the SR Ca²⁺ channel. The II-III loop model of EC coupling is simple, has intuitive appeal, and has received broad consideration. However, in reality, the signaling mechanism may be more complex. The prevailing evidence indicates that interactions between 1S and RyR1 are likely to involve the II-III loop, but also many other domains of 1S (7,19–24). Additional complexity is brought about by a deletion analysis showing that the II-III loop may not account entirely for the signaling function of the DHPR (25). Furthermore, the DHPR ß1a subunit is essential for skeletal-type EC coupling (26–28), and interactions between this subunit and RyR1 are almost certain (6,29). Hence, the molecular determinants of the voltage-dependent mechanism by which the DHPR activates RyR1 may be broader than initially anticipated by the II-III loop model and are still open to debate despite unrelenting efforts.

In light of the growing multiplicity of DHPR-RyR1 interactions, here we reinvestigated the contribution of the cytosolic domains of 1S to skeletal-type EC coupling in dysgenic (1S null) myotubes. Previous studies have focused almost exclusively on the functional identity of regions within the skeletal II-III loop (14,16,17,25,30,31). However, a voltage-clamp analysis of the EC-coupling phenotype contributed by each of the 1S interrepeat loops, as well as by the N- and C-terminal domains, has not been previously conducted. Likewise, the phenotype contributed by all the cytosolic domains together, as they would be present in the intact subunit, has not been documented. Since 1C does not express skeletal-type EC coupling (14,32), we focused on chimeras consisting of 1C (Ca_V1.2) with sequences from 1S (Ca_V1.1), substituting the interrepeat loops (I-II, II-III, or III-IV) or the N- or C-terminus . We report that Ca²⁺ entry-independent Ca²⁺ transients of a magnitude and voltage dependence similar to those expressed by wild-type (WT) 1S required a chimera with multiple 1S domains, including the II-III loop. However, the chimera with the II-III loop alone was insufficient. Some of these results have been published in abstract form (33,34).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Identification of genotypes
Dysgenic (1S-null mdg) mice were screened for both wild-type and mutant alleles of the DHPR 1S gene. Digestion of tail samples and subsequent verification of genotypes by the polymerase chain reaction (PCR) were described previously (27).

Primary cultures and cDNA transfection
Cultures of myotubes were prepared from hind limbs of E18 fetuses, as described previously (35). Cultures were grown at 37°C in 8% CO₂ gas. After myoblast fusion (5 days), the medium was replaced with FBS-free medium, and CO₂ was decreased to 5%. cDNA transfection was performed during the myoblast fusion stage with the polyamine LT1 (Panvera, Madison, WI). 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 anti-CD8 antibody (Dynal ASA, Oslo, Norway). Whole-cell analysis of Ca²⁺ currents, charge movements, and Ca²⁺ transients was performed 3–5 days post-transfection. The numbers of separate myotube cultures that were transfected and from which data were collected were as follows for Ca²⁺ current, Ca²⁺ transient, and charge movements when applicable: for nontransfected mdg, 10, 27, and 2; for WT 1S, 7, 3, and none; for 1S(E1014K), 3, 4, and 2; for WT 1C, 5, 3, and none; for 1C(E736K), 3, 2, and 2; for 1C/1S N, 3, 3, and none; for 1C/1S I-II, 4, 4, and none; for 1C/1S II-III, 6, 4, and none; for 1C(E736K)/1S II-III, 3, 2, and 3; for 1C/1S III-IV, 4, 3, and none; for 1C/1S C, 3, 2, and none; for 1C/1S all loop, 3, 3, and none; for 1C(E736K)/1S all loop: 3, 3, and 3; for all cadmium experiments, 2, 2, and none.

cDNA constructs
Chimeric variants were made by two-step PCR strategies using cDNAs for rabbit skeletal muscle 1S (residues 1–1873; Genbank No. M23919) and rabbit cardiac 1C (residues 1–2171; Genbank No. X15539). The PCR products were subcloned into pCR-Blunt vector (Invitrogen, Carlsbad, CA), excised by digestion with AgeI/NotI, and cloned into the pSG5 vector (Stratagene, San Diego, CA) in frame with the first 11 residues of the phage T7 gene 10 protein for antibody tagging. All constructs were sequenced twice or more at a campus facility.

Domain boundaries and nomenclature
Alignment of 1S and 1C sequences was performed with DNASTAR (Madison, WI) using the Jotun-Hein method. 1S N corresponds to residues 1–51 and replaced 1C N residues 1–154; 1S I-II loop corresponds to residues 335–432 and replaced 1C I-II loop residues 436–554; 1S II-III loop corresponds to residues 667–799 and replaced 1C II-III loop residues 789–930; 1S III-IV loop corresponds to residues 1067–1120 and replaced 1C III-IV loop residues 1188–1241; 1S C corresponds to residues 1328–1873 and replaced 1C C residues 1507–2171.

1C/1S N
This chimera consists of 1S residues M1–K51 fused to the N-terminus of 1C residues P155–L2171. The N-terminus of 1S was amplified from full-length 1S and corresponds to residues 1–51. The second-step PCR product containing the 1S N-terminus and part of 1C domain I was fused to the pSG5 1C vector using NheI/SacI sites.

1C/1S I-II
This chimera consists of 1C residues M1–S435 fused to the N-terminus of 1S residues G335–R432 fused to the N-terminus of 1C residues V555–L2171. The I-II loop of 1S was amplified from full-length 1S and corresponds to residues 335–432. The second-step PCR product containing the 1S I-II loop and part of 1C domains I and II was fused to the pSG5 1C vector using BamHI/XhoI sites.

1C/1S II-III
This chimera consists of 1C residues M1–D788 fused to the N-terminus of 1S residues A667–T799 fused to the N-terminus of 1C residues I931–L2171. To replace the II-III loop, a HindIII site at nucleotide 2561 and a SpeI site at nucleotide 3203 were introduced into full-length 1C as silent mutations. The II-III loop of 1S was amplified from full-length 1S, and corresponds to residues 667–799. The second-step PCR product containing the 1S loop and part of 1C domain III was fused to the pSG5 1C vector using HindIII/SpeI sites.

1C/1S III-IV
This chimera consists of 1C residues M1–V1187 fused to the N-terminus of 1S residues T1067–F1120 fused to the N-terminus of 1C residues E1242–L2171. The III-IV loop of 1S was amplified from full-length 1S and corresponds to residues 1067–1120. The second step PCR product containing the 1S III-IV loop and part of 1C domains III and IV was fused to the pSG5 1C vector using SpeI/SacII sites.

1C/1S C
This chimera consists of 1C residues M1–M1506 fused to the N-terminus of 1S residues D1328–P1873. The C-terminus of 1S was amplified from full-length 1S using a forward primer containing a 5' overhang of the C-terminal end of 1C domain IV up to the BclI restriction site, and a reverse primer at the stop codon of 1S. The PCR product containing the 1S C-terminus and part of 1C domain IV was fused to the pSG5 1C vector using BclI/NotI sites.

1C/1S all loop (N, I-II, II-III, III-IV, C)
This chimera consists of fusions of the following peptide fragments in sequential order from N- to C-terminus: 1S(M1–K51)/1C(P155–S435)/1S(G335–R432)/1C(V555–D788)/1S(A667–T799)/1C(I931–V1187)/1S(T1067–F1120)/1C(E1242–M1506)/1S(D1382–P1873). This chimera was made by a cut-and-paste method using the chimeras and restriction sites indicated above.

1C(E736K)
This domain II pore mutation was described elsewhere (36) and consists of a replacement of the glutamate residue at position 736 by lysine. We designed a 37-base antisense primer that introduced a mismatch at nucleotide 2397 (g2397a), and a sense primer that annealed before the BamHI site of 1C. The PCR product was cloned into the pSG5 1C vector at BamHI/EcoRI sites.

1S(E1014K)
This domain III pore mutation consists of replacement of a glutamate residue at position 1014 by lysine, and was previously made and described elsewhere (25).

Whole-cell voltage clamp
Whole-cell recordings were performed 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. Patch pipettes had a resistance of 1–3 M when filled with the pipette solution. To obtain Ca²⁺ conductance curves, cells were maintained at a holding potential of –40 mV, and depolarized in ascending order every 3 s. The pulse duration was 500 ms and was changed in 5-mV increments up to +85 mV. To obtain Ca²⁺ transient curves, cells were maintained at –40 mV and depolarized in descending order every 30 s to permit recovery of the resting fluorescence. The pulse duration was 200 ms and was changed in 20-mV decrements from +90 mV to –30 mV. To obtain charge movement curves, we used a P/4 protocol with a long prepulse to inactivate Na⁺-channel ionic and gating currents (25). The pulse protocol was as follows. The command voltage was stepped from a holding potential of –80 mV to –30 mV for 698 ms, to –50 mV for 5 ms, to the test potential for 50 ms, to –50 mV for 50 ms, and then to the –80 mV holding potential. Test potentials were applied in decreasing order every 10 mV from +100 to –80 mV. The waiting period between test pulses was 10 s.

Confocal fluorescence microscopy
Ca²⁺ transients were measured by confocal line scanning at room temperature. Cells were loaded with 5 µM fluo-4 acetoxymethyl ester (Molecular Probes, Eugene, OR) for 1 h at room temperature. Cells were viewed with an inverted IX20 Olympus microscope with a 20x (NA 1.4) objective and a Fluoview confocal attachment (Olympus, Melville, NY). The 488 nm line provided by a 5 mW Argon laser was attenuated to 6% with neutral density filters. Line scans were acquired at a speed of 2.05 ms per line. All line scans consisted of 1000 lines at a width of 512 pixels. The spatial dimension of the line scan was 30–60 µm , and covered the entire width of the myotube. Locations selected for line scans were devoid of nuclei and had a low resting fluorescence. Line scans were synchronized to start 100 ms before the onset of the depolarization for voltage-clamp experiments. The time course of the space-averaged fluorescence intensity change was estimated as described elsewhere (26,27) and is reported in F/F units. The peak-to-peak noise in the baseline fluorescence averaged 0.1 F/F units. Image analyses were performed with NIH Image software (National Institutes of Health, Bethesda, MD).

Solutions
For Ca²⁺ currents and Ca²⁺ transients, the external solution was (in mM) 130 TEA methanesulfonate, 10 CaCl₂, 1 MgCl₂, 10^–3 TTX, and 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. For charge movements, the internal solution was (in mM) 120 NMG (N-methyl glucamine)-Glutamate, 10 HEPES-NMG, and 10 EGTA-NMG, pH 7.3. This solution produced a more reliable block of the outward ionic current than the internal solution used for Ca²⁺ currents. The external solution was supplemented with 0.5 mM CdCl₂ to block background Ca²⁺ currents present in mdg myotubes, 0.5 mM LaCl₃ to increase pipette seal resistance and 0.05 mM TTX to block residual Na⁺ current.

Curve fitting
The voltage dependence of the Ca²⁺ conductance, charge movements, and sigmoidal fluorescence-voltage relationships were fitted with a standard Boltzmann equation:

(1)

where V (in mV) is the test potential, A_max is G_max, Q_max, or F/F_max, V_1/2 (in mV) is the midpoint potential, and k (in mV) is the slope factor. For bell-shaped fluorescence-voltage curves, the Boltzmann equation was modified as follows:

(2)

where Vr (in mV) is a constant that accounts for the decrease in Ca²⁺ transient amplitude at positive potentials, and k' (in mV) is an empirical scaling factor (26,27). Other parameters are the same as in Eq. 1. Parameters of a fit of averages of many cells (population average) are shown in the figures. Parameters of the fit of individual cells are shown in tables. Parameters of the fit of the population average differed slightly from the mean of the fit of individual cells. The latter parameters generated theoretical curves that were a better fit with the average data and, for that reason, were used in the figures. Analysis of variance (ANOVA) was performed with Analyze-it (Leeds, UK).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Recovery of voltage- or Ca²⁺ entry-dependent EC coupling in dysgenic myotubes
Studies have shown that 1C is targeted to EC-coupling junctions in cultured skeletal myotubes, and that the Ca²⁺ current generated by 1C can trigger Ca²⁺ transients by Ca²⁺-dependent activation of RyR1 (31,32). Chimeras with 1C in the backbone also activate Ca²⁺ transients by a similar mechanism when expressed in skeletal myotubes (11,14). Hence, Ca²⁺ release induced by the Ca²⁺ current is inherent to chimeras of 1C, and this component needs to be carefully separated from the release of interest triggered by mechanical DHPR-RyR1 coupling. To exclude the component of the Ca²⁺ transient triggered by the Ca²⁺ current without changing the external solution, we mutated the conserved glutamate E736 in the P-loop of repeat II of 1C, previously shown to be critical for Ca²⁺ permeation (36). Fig. 1 shows the EC coupling and Ca²⁺ current characteristics of 1C(E736K) and 1S(E1014K), localized in the P-loop of repeat III of 1S (25,37). The top traces show representative Ca²⁺ transients and Ca²⁺ currents at +30 mV, with the pore mutants depicted by shaded traces. In this and other figures, the displayed Ca²⁺ transients and Ca²⁺ currents were obtained with different protocols and different internal solutions (see Materials and Methods). For Ca²⁺ currents, we used 500-ms depolarizations in a highly buffered internal solution (5 mM EGTA). These conditions permitted the determination of the time course of inactivation while maintaining a low cytosolic Ca²⁺ at all potentials. For Ca²⁺ transients, we used 200-ms depolarizations in a lightly buffered internal Ca²⁺ solution (0.1 mM EGTA). These conditions limited SR Ca²⁺ release, increased the rate of resting Ca²⁺ recovery, and increased the intensity of the fluorescence signal. In all cases, the external Ca²⁺ concentration was 10 mM. WT 1S expressed slow-activating/noninactivating Ca²⁺ currents typical of cultured normal primary myotubes (35). In contrast, WT 1C expressed fast-activating/slow-inactivating Ca²⁺ currents, in agreement with previous results in mdg myotubes (11,32,38). The degree of inactivation of 1C was variable, and to some extent varied with the Ca²⁺ current density, consistent with Ca²⁺-entry-dependent inactivation of 1C investigated by others in myotubes (16). Both 1C and 1S recovered a Ca²⁺ current density close to that of normal myotubes reported elsewhere (35). Inward currents expressed by the pore mutants were drastically reduced. In the case of 1S(E1014K), inward current was undetectable (<0.1 pA/pF), consistent with previous results (25). In the case of 1C(E736K), the inward current at +30 mV was reduced 180-fold compared to WT 1C, consistent with determinations in oocytes (36). The reduction was less pronounced if calculations are based on the maximum Ca²⁺ conductance, G_max, expressed by the conducting and nonconducting variants (see Tables 1 and 2). This is because the G_max of 1S(E1014K) and 1C(E736K) is dominated by the conductance of the outward current (Fig. 1, middle panels). The latter is a mixture of outward current through the mutant Ca²⁺ channel and background outward currents unaffected by the mutation.

View larger version (23K): [in this window] [in a new window]   FIGURE 1  Ca2+ currents and Ca2+ transients expressed by 1S and 1C variants. Columns show representative mdg myotubes transfected with WT 1S, WT 1C, and variants carrying pore mutations, namely 1S (E1014K) and 1C (E736K). Shaded traces and shaded symbols correspond to the pore mutant form of the variant indicated above each column. The top trace corresponds to the spatial integral of the confocal Ca2+ transient in F/F units in response to a 200-ms depolarization to +30 mV from a holding potential of –40 mV. The second row of traces corresponds to the Ca2+ current elicited during a 500-ms depolarization from a holding potential of –40 mV to +30 mV. The Ca2+ transients and Ca2+ currents shown in this and other figures were obtained with different protocols (see text, especially Materials and Methods). Population averages of the Ca2+ current-voltage curves are located centrally. These were obtained by depolarizing transfected myotubes in 5-mV increments from a holding potential of –40 mV for 500 ms. At the bottom are Ca2+ transient-voltage curves for population averages in response to a 200-ms depolarization from –40 mV to the indicated potentials. The population averaged fluorescence-voltage curves for 1S and 1S(E1014K) were fit with Eq. 1, and those obtained in myotubes expressing 1C and 1C(E736K) were fit with Eq. 2. Curves were fit with the following parameters (F/Fmax in F/F units, V1/2 in mV, and k in mV, respectively): for 1S: 2.6, 5.3, and 11.7; for 1S(E1014K), 2.6, 6.4, and 7.8; for 1C, 1.2, 20.6, and 6.1; for 1C(E736K), 0, –19.7, and 20. For a fit of parameters by cell and number of cells included in the fit, see Table 1.

View this table: [in this window] [in a new window]   TABLE 1  Ca2+ conductance and Ca2+ transients expressed by 1C/1S chimeras in dysgenic (mdg) skeletal myotubes

Incomplete recovery of skeletal-type EC coupling by the 1S II-III loop revealed by pore mutations that eliminate inward Ca²⁺ current
Ca²⁺ currents and transients expressed by chimeras consisting of 1C carrying each of the cytosolic domains of 1S, namely the N-terminal, I-II loop, II-III loop, III-IV loop, or C-terminal domains, are shown in Fig. 2. Coordinates of skeletal domains donated to 1C, and of homologous regions removed from 1C, are described in Materials and Methods. The top traces correspond to inward Ca²⁺ currents for depolarizations to –30 mV and +30 mV in dysgenic myotubes expressing each of the five indicated chimeras. All chimeras expressed high-density, fast-activating, and slow-inactivating Ca²⁺ currents typical of WT 1C when expressed in myotubes. Furthermore, the fitted G_max was statistically similar to that of 1C or 1S (Table 1). Two chimeras, that carrying the N-terminal domain (labeled 1C N) and that carrying the II-III loop (labeled 1C/1S II-III), expressed Ca²⁺ current at a density higher than that of WT 1C, although the difference was not significant. Representative Ca²⁺ transients at +30 mV, and the voltage dependence of the Ca²⁺ transient amplitude measured at the end of a 200-ms depolarization, are shown in the bottom graphs of Fig. 2. For reference, the shaded lines in the plots indicate the fitted fluorescence-voltage relationship of myotubes expressing WT 1S and WT 1C described above. The II-III loop chimera consistently expressed Ca²⁺ transients that were much larger in amplitude than those expressed by the other chimeras. Furthermore, only the II-III loop chimera expressed a fluorescence-voltage relationship that was sigmoidal like that of WT 1S. All other chimeras expressed fluorescence-voltage relationships that were bell-shaped like that of WT 1C, indicative of a Ca²⁺-dependent mechanism. The bell-shaped voltage dependence is particularly prominent for the III-IV loop and the C-terminal domain chimeras. Furthermore, in these cases there was a significant negative shift in midpoint potential compared to WT 1C. The shifts were consistent with differences in midpoint potential found for the Ca²⁺ conductance described in Table 1. In summary, only the 1C/1S II-III loop chimera expressed Ca²⁺ transients with a sigmoidal fluorescence-voltage curve indicative of skeletal-type EC coupling. This result is consistent with previous observations, which have suggested that the II-III loop is a critical domain for skeletal-type EC coupling (11,13–17,30,32). Other domains of 1S, namely the III-IV loop and the C-terminal domain, affected channel gating by shifting Ca²⁺ current activation to more negative potentials, resulting in a concurrent shift in the voltage dependence of the Ca²⁺ transient. Furthermore, the skeletal N-terminal domain and the I-II loop do not appear to influence the Ca²⁺-dependent EC-coupling phenotype in a significant manner.

View larger version (25K): [in this window] [in a new window]   FIGURE 2  Ca2+ currents and Ca2+ transients expressed by 1C/1S chimeras. Columns show representative mdg myotubes transfected with the identified 1C/1S chimeras. Nomenclature and domain boundaries for the chimeras are described in Materials and Methods. The top traces correspond to Ca2+ currents from the same cell in response to a 500-ms depolarization from a holding potential of –40 mV to –30 mV (no current) and to +30 mV (near maximum inward current). Population averages of the Ca2+ current-voltage curves are located centrally. These were obtained by depolarizing transfected myotubes in 5-mV increments from a holding potential of –40 mV for 500 ms. The traces in the middle correspond to confocal Ca2+ transients in mdg myotubes expressing each of the chimeras in F/F units in response to a 200-ms depolarization to +30 mV from a holding potential of –40 mV. At the bottom are Ca2+ transient-voltage curves for population averages in response to a 200-ms depolarization from –40 mV to the indicated potentials. The shaded traces without data in each graph correspond to the fitted voltage dependence of WT 1S and WT 1C from Fig. 1. The fluorescence-voltage curves for all 1C/1S chimeras were fit with Eq. 2, except that obtained in myotubes expressing 1C/1S II-III, which was fit with Eq. 1. Curves (black) were fit with the following parameters (F/Fmax in F/F units, V1/2 in mV, and k in mV, respectively): for 1C/1S N, 0.5, 16.7, and 10.0; for 1C/1S I-II, 1.3, 20.0, and 9.9; for 1C/1S II-III, 2.4, 2.8, and 4.9. For 1C/1S III-IV, 1.0, 1.2, and 4.5; and for 1C/1S C, 0.8, 4.9, and 3.9.

View larger version (26K): [in this window] [in a new window]   FIGURE 3  Full restoration of skeletal-type EC coupling by the all loop chimera and partial restoration by the II-III loop chimera. Columns show representative mdg myotubes transfected with the 1C/1S II-III loop chimera and 1C/1S all loop chimera, both in the presence and absence of the pore mutation E736K. Shaded traces and shaded symbols correspond to pore mutants. The top traces correspond to the spatial integral of the confocal Ca2+ transient in F/F units in response to a 200-ms depolarization to +30 mV from a holding potential of –40 mV. The second row of traces corresponds to the Ca2+ current elicited during a 500-ms depolarization from –40 mV to +30 mV. Population averages of the Ca2+ current-voltage curves are located centrally. These were obtained by depolarizing transfected myotubes in 5-mV increments from a holding potential of –40 mV for 500 ms. At the bottom are Ca2+ transient-voltage curves for population averages in response to a 200-ms depolarization from –40 mV to the indicated potentials. All fluorescence-voltage curves were fit with Eq. 1. Curves were fit with the following parameters (F/Fmax in F/F units, V1/2 in mV, and k in mV, respectively): for 1C/1S II-III, 2.4, 2.8, and 4.9; for 1C(E736K)/1S II-III, 0.9, 17.1, and 15.5; for 1C/1S all loop, 2.1, 1.7, and 10.9; for 1C(E736K)/1S all loop: 2.1, 14.3, and 8.

Fig. 4 shows normalized charge movement density (fC/pF) expressed by the II-III loop chimera, the all loop chimera, and the two WT constructs. We carried out a standard gating current protocol (25) in myotubes expressing variants carrying the pore mutation. This reduced the ionic current, and hence increased the success of the measurement. The traces in the insets show the transient outward ON current at the start of a 50-ms test voltage step to +60 mV followed by inward OFF current at the end of the voltage step as charges return to the original position. The complete pulse protocol (see Materials and Methods) included a 698-ms prepulse depolarization to –30 mV to remove immobilization-sensitive components of the gating current unrelated to the DHPR, and pretest P/4 pulses to eliminate linear components unrelated to voltage sensors. The graphs show normalized charge density-voltage relationships obtained by integration of the OFF component which, in our hands, was less contaminated by residual ionic current than the ON component. The main contaminant of the ON component was an 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 –40 mV and saturated at potentials more positive than +60 mV. Data were adequately fit by a Boltzmann equation (Eq. 1). In nontransfected myotubes, we detected a small background charge with a maximum density of 1 fC/pF, similar to that reported previously (25). The average charge movement density of nontransfected cells is shown by the line without data, and the parameters of the fit are shown in Table 2. The maximum charge-movement densities expressed by the 1C(E736K)/1S II-III loop chimera, the 1C(E736K)/1S all loop chimera, 1C(E736K), and 1S(E1014K) were statistically similar (Table 2). These values agreed with previous determinations in dysgenic myotubes expressing WT 1C and WT 1S (12,25). The results show that DHPR voltage-sensor density was not a confounding factor in our studies .

View larger version (21K): [in this window] [in a new window]   FIGURE 4  Charge movement density expressed 1C/1S chimeras carrying pore mutations. Graphs show the population-averaged voltage dependence of the OFF charge in femto-Coulombs normalized to the linear membrane capacitance of each cell in pico-Farads. The OFF charge was obtained by integration of the OFF charge movement current at the end of a 50-ms test pulse after on-line subtraction of the linear component. Measurements were made in mdg myotubes transfected with the indicated constructs using a pulse protocol and external and pipette solutions described in Materials and Methods. The shaded trace in each graph corresponds to the fit of the population averaged charge movement of nontransfected dysgenic myotubes. Insets are representative traces of charge movement currents during the test-pulse phase of the pulse protocol elicited from a holding potential of –80 mV to a test potential of +60 mV for 50 ms. Charge movement versus voltage curves were fit with Eq. 1 with the following parameters (Qmax in fC/pF, V1/2 in mV, and k in mV, respectively); for 1S(E1014K), 5.3, 8.2, and 19.1; for 1C(E736K), 5.9, 2.7, and 17.6; for 1C(E736K)/1S II-III, 5.7, 10.6, and 20.1; for 1C(E736K)/1S all loop, 4.4, 12.0, and 17.6; and for nontransfected dysgenic myotubes (shaded traces): 1.4, –7.0, and 12.9. For a fit of parameters by cell and number of cells included in the fit, see Table 2.

Incomplete recovery of skeletal-type EC coupling by the 1S II-III loop revealed by Cd²⁺ block of inward Ca²⁺ current

View larger version (31K): [in this window] [in a new window]   FIGURE 5  Cd2+ block of Ca2+-dependent EC coupling promoted by the II-III loop chimera. Columns show representative mdg myotubes expressing the II-III loop and all loop chimeras. The left panel shows myotubes expressing WT 1S used as control. The top graphs correspond to population average current-voltage curves for depolarizations of 500 ms from a holding potential of –40 mV. Insets show representative Ca2+ currents elicited at +30 mV from a holding potential of –40 mV. With the exception of the bar graphs at the bottom, all shaded traces and shaded symbols are in 10–4 M CdCl2 added to the bath solution. The middle graphs show population-averaged Ca2+ transient-voltage relationships for depolarizations of 200 ms from a holding potential of –40 mV. Insets correspond to the spatial integral of the confocal fluorescence in F/F units in response to a 200-ms depolarization to +30 mV from a holding potential of –40 mV. Fluorescence-voltage curves were fit with Eq. 1. Curves were fit with the following parameters (Fmax in F/F units, V1/2 in mV, and k in mV, respectively): for WT 1S in 0 Cd2+, 2.5, 13.7, and 14.1; for WT 1S in 10–4 Cd2+, 2.2, 3.0, and 8.2; for 1C/1S II-III in 0 Cd2+, 2.8, 2.8, and 4.7; for 1C/1S II-III in 10–4 Cd2+, 0.7, 17.0, and 13.5; for 1C/1S all loop in 0 Cd2+, 2.1, 1.7, and 10.9; for 1C/1S all loop in 10–4 Cd2+, 2.7, 15.6, and 9.3. For a fit of parameters by cell and number of cells included in the fit, see Table 3. The bottom bar graphs correspond to normalized Ca2+ currents (shaded bars) and normalized F/Fmax (black) in control solution without Cd2+, external solution with 10–6 M CdCl2, and external solution with 10–4 M CdCl2. Ca2+ currents at +30 mV were normalized to the population averaged Ca2+ current at +30 mV in the absence of Cd2+. F/Fmax from a fit of individual cells was normalized to F/Fmax from a fit of the population average in the absence of Cd2+.

View this table: [in this window] [in a new window]   TABLE 3  Ca2+ conductance and Ca2+ transients expressed by 1C/1S chimeras in presence of extracellular cadmium

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

The Ca²⁺ transient expressed by CSk3, the 1C/1S II-III loop chimera described by Tanabe et al. (11) was substantially reduced in the presence of inorganic blockers of the inward Ca²⁺ current. Ca²⁺ fluorescence-voltage relationships for CSk3 have not been described (11,14); therefore, it is difficult to compare the two sets of results. However, qualitatively the results are similar and indicate that both II-III loop chimeras have a sizable EC-coupling component unrelated to the skeletal phenotype. CSk3 consists of 1C(1–787)/1S(666–791)/1C(923–2171), whereas our chimera included eight additional residues, namely 1S(1–788)/1S(667–799)/1C(931–2171). These differences may not be significant since critical determinants within the 1S II-III loop have been mapped to the segment of residues 671–690 and 720–765 present in both chimeras (14,25). Furthermore, charge movements expressed by both chimeras in dysgenic myotubes are comparable (12) (Table 2). Hence, possible differences in voltage-sensor densities, which could conceivably affect the density of mechanically coupled RyR1 channels, are nonexistent. Interestingly, Ca²⁺ fluorescence-voltage relationships have been published for GFP-CSk53, an 1C/1S chimera carrying the 1S II-III loop residues 720–765 deemed critical for skeletal-type EC coupling. This chimera also has an N-terminal fusion of the green fluorescent protein (GFP). The F_max expressed by GFP-CSk53 was considerably lower than that expressed by the GFP-WT 1S fusion even in external solution containing the standard 10 mM Ca²⁺ (31). Thus, the low F_max of a II-III loop chimera may not be related in all instances to the presence of an enlarged Ca²⁺-dependent EC-coupling component, which is removed when the inward Ca²⁺ current is blocked. It could be that in this case, the GFP moiety hinders EC coupling in the CSk53 chimera, or that residues 720–765 are insufficient for a quantitative EC-coupling recovery comparable to that of WT 1S. In support of the latter, it is important to note that the identification of critical 1S II-III loop residues 720–765 in chimeras without GFP was achieved at the expense of a substantial loss in Ca²⁺ transient amplitude relative to the Ca²⁺ transient expressed by the parent CSk3 chimera carrying the complete 1S II-III loop (14). Thus, an appropriate structural context is essential for a quantitative recovery of EC-coupling function.

It was intriguing to find that, on average, the shape of the fluorescence-voltage curve for the Ca²⁺-conducting and pore-mutant variants of the 1C/1S II-III loop chimera were both sigmoidal (Fig. 3). A somewhat bell-shaped curve was expected for the former based on the significant dependence on the Ca²⁺ current for triggering release. We considered the possibility that the reduction in F/F_max observed in the pore-mutant variant of the II-III loop chimera reflected inactivation of the voltage sensor due to the elimination of a critical C