The II-III Loop of the Skeletal Muscle Dihydropyridine Receptor Is Responsible for the Bi-directional Coupling with the Ryanodine Receptor
1999; Elsevier BV; Volume: 274; Issue: 31 Linguagem: Inglês
10.1074/jbc.274.31.21913
ISSN1083-351X
AutoresManfred Grabner, Robert T. Dirksen, Norio Suda, Kurt G. Beam,
Tópico(s)Neurobiology and Insect Physiology Research
ResumoThe dihydropyridine receptor (DHPR) in the skeletal muscle plasmalemma functions as both voltage-gated Ca2+ channel and voltage sensor for excitation-contraction (EC) coupling. As voltage sensor, the DHPR regulates intracellular Ca2+ release via the skeletal isoform of the ryanodine receptor (RyR-1). Interaction with RyR-1 also feeds back to increase the Ca2+ current mediated by the DHPR. To identify regions of the DHPR important for receiving this signal from RyR-1, we expressed in dysgenic myotubes a chimera (SkLC) having skeletal (Sk) DHPR sequence except for a cardiac (C) II-III loop (L). Tagging with green fluorescent protein (GFP) enabled identification of expressing myotubes. Dysgenic myotubes expressing GFP-SkLC or SkLC lacked EC coupling and had very small Ca2+currents. Introducing a short skeletal segment (α1Sresidues 720–765) into the cardiac II-III loop (replacing α1C residues 851–896) of GFP-SkLC restored both EC coupling and Ca2+ current densities like those of the wild type skeletal DHPR. This 46-amino acid stretch of skeletal sequence was recently shown to be capable of transferring strong, skeletal-type EC coupling to an otherwise cardiac DHPR (Nakai, J., Tanabe, T., Konno, T., Adams, B., and Beam, K.G. (1998) J. Biol. Chem.273, 24983–24986). Thus, this segment of the skeletal II-III loop contains a motif required for both skeletal-type EC coupling and RyR-1-mediated enhancement of Ca2+ current. The dihydropyridine receptor (DHPR) in the skeletal muscle plasmalemma functions as both voltage-gated Ca2+ channel and voltage sensor for excitation-contraction (EC) coupling. As voltage sensor, the DHPR regulates intracellular Ca2+ release via the skeletal isoform of the ryanodine receptor (RyR-1). Interaction with RyR-1 also feeds back to increase the Ca2+ current mediated by the DHPR. To identify regions of the DHPR important for receiving this signal from RyR-1, we expressed in dysgenic myotubes a chimera (SkLC) having skeletal (Sk) DHPR sequence except for a cardiac (C) II-III loop (L). Tagging with green fluorescent protein (GFP) enabled identification of expressing myotubes. Dysgenic myotubes expressing GFP-SkLC or SkLC lacked EC coupling and had very small Ca2+currents. Introducing a short skeletal segment (α1Sresidues 720–765) into the cardiac II-III loop (replacing α1C residues 851–896) of GFP-SkLC restored both EC coupling and Ca2+ current densities like those of the wild type skeletal DHPR. This 46-amino acid stretch of skeletal sequence was recently shown to be capable of transferring strong, skeletal-type EC coupling to an otherwise cardiac DHPR (Nakai, J., Tanabe, T., Konno, T., Adams, B., and Beam, K.G. (1998) J. Biol. Chem.273, 24983–24986). Thus, this segment of the skeletal II-III loop contains a motif required for both skeletal-type EC coupling and RyR-1-mediated enhancement of Ca2+ current. Excitation-contraction (EC) 1The abbreviations used are: The abbreviations used are: EC, excitation-contraction; DHPR, dihydropyridine receptor; RyR-1, skeletal ryanodine receptor; GFP, green fluorescent protein; SR, sarcoplasmic reticulum; nt, nucleotide number; V, ohms; F, farads; S, siemens; C, coulombs.. coupling in skeletal muscle depends upon a functional interaction between dihydropyridine receptors (DHPRs) in the plasmalemma and ryanodine receptors (RyRs) in the sarcoplasmic reticulum (SR). In skeletal muscle, the DHPR functions both as an L-type Ca2+ channel and as the voltage sensor, which in response to plasmalemmal depolarization, transmits a signal that causes RyR-1 (the skeletal RyR isoform) to release Ca2+ from the SR (1Rios E. Brum G. Nature. 1987; 325: 717-720Crossref PubMed Scopus (654) Google Scholar, 2Tanabe T. Beam K.G. Powell J.A. Numa S. Nature. 1988; 336: 134-139Crossref PubMed Scopus (584) Google Scholar, 3Takeshima H. Iino M. Takekura H. Nishi M. Kuno J. Minowa O. Takano H. Noda T. Nature. 1994; 369: 556-559Crossref PubMed Scopus (326) Google Scholar). The nature of the signal transmitted from the skeletal DHPR to RyR-1 is not yet understood, although there is strong evidence that skeletal-type EC coupling does not rely upon the entry of external Ca2+ (4Armstrong C.M. Bezanilla F.M. Horowicz P. Biochim. Biophys. Acta. 1972; 267: 605-608Crossref PubMed Scopus (301) Google Scholar). An approach to identifying regions of the skeletal DHPR that are important for EC coupling has been to express cDNAs encoding chimeric DHPRs in dysgenic myotubes, which lack endogenous skeletal DHPRs. This work has shown that a purely cardiac DHPR expressed in dysgenic myotubes results in EC coupling which is cardiac type (dependent on entry of Ca2+) (5Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 344: 451-453Crossref PubMed Scopus (194) Google Scholar), whereas skeletal-type EC coupling results from expression of a chimeric DHPR having cardiac sequence except for a skeletal II-III loop (6Tanabe T. Beam K.G. Adams B.A. Niidome T. Numa S. Nature. 1990; 346: 567-569Crossref PubMed Scopus (493) Google Scholar). More recently, it was shown that strong skeletal-type coupling could be produced by a chimeric DHPR that contained only a 46-amino acid skeletal segment within the II-III loop and weak skeletal-type coupling by a chimera containing only an 18-amino acid skeletal segment (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar). Analysis of myotubes from dyspedic mice, which lack RyR-1, has revealed that in addition to the orthograde (EC coupling) signal transmitted from the skeletal DHPR to RyR-1, there also appears to be a retrograde signal whereby RyR-1 increases the magnitude of the L-type Ca2+ current mediated by the DHPR. In particular, Ca2+ current density is very low in dyspedic myotubes even though the surface density of DHPRs appears to be essentially normal (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar, 9Fleig A. Takeshima H. Penner R. J. Physiol. (Lond.). 1996; 496: 339-345Crossref Scopus (29) Google Scholar). Expression of cDNA encoding RyR-1 causes the density of L-type current in dyspedic myotubes to increase toward normal (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar). However, these experiments did not reveal whether the region of the skeletal DHPR that is crucial for orthograde coupling is also important for the RyR-1-mediated enhancement of DHPR Ca2+current. Here we describe experiments to identify regions of the skeletal DHPR that are critical for the ability of the DHPR to receive the retrograde (current-enhancing) signal from RyR-1. The results demonstrate that the II-III loop is critical for both orthograde and retrograde signaling. Within the II-III loop, the 46-amino acid segment found to be important for skeletal-type EC coupling is also important for transducing the retrograde signal from RyR-1. Chimeras between the α1 subunits of the skeletal muscle DHPR (Sk (10Tanabe T. Takeshima H. Mikami A. Flockerzi V. Takahashi H. Kangawa K. Kojima M. Matsuo H. Hirose T. Numa S. Nature. 1987; 328: 313-318Crossref PubMed Scopus (970) Google Scholar)) and the cardiac muscle DHPR (C (11Mikami A. Imoto K. Tanabe T. Niidome T. Mori Y. Takeshima H. Narumiya S. Numa S. Nature. 1989; 340: 230-233Crossref PubMed Scopus (770) Google Scholar)) had amino acid composition (numbers in parentheses) as follows. SkLC: Sk (1–654), C (777–927), Sk (797–1873). SkLCS46: Sk (1–654), C (777–850), Sk (720–765), C (897–927), Sk (797–1873). SkLCS18: Sk (1–654), C (777–855), Sk (725–742), C (874–927), Sk (797–1873). The chimeras were constructed and inserted into mammalian expression vectors as described below (nucleotide numbers (nt) indicated in parentheses): The EcoRI-XmnI fragment of Sk (nt 1007–1964) was coligated with the ligation product from theXmnI-HincII fragment of C (nt 2330–2782) plus the HincII-XhoI fragment of Sk (nt 2389–2654) into the corresponding EcoRI/XhoI restriction sites of a SacII-XhoI subclone of Sk (nt 86–2654) in pBluescript SK+ (Stratagene). Finally, theSacII-XhoI insert of the modified subclone (now carrying the cardiac II-III loop sequence) was ligated into the corresponding SacII/XhoI restriction sites of plasmid pCAC6, which contains the complete skeletal DHPR coding region in the mammalian expression vector pKCRH2 (6Tanabe T. Beam K.G. Adams B.A. Niidome T. Numa S. Nature. 1990; 346: 567-569Crossref PubMed Scopus (493) Google Scholar). The EcoRI-XmnI fragment of Sk (nt 1007–1964) was coligated with theXmnI-AflII* fragment (nt C2330-Sk2297) (partial cut) of clone CSk53 (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar) which is C (1–850), Sk (720–765), C (897–2171) into the corresponding EcoRI/AflII restriction sites (nt Sk1007/C2690) of the modified (the cardiac II-III loop carrying) SacII-XhoI subclone of Sk (nt 86–2654) in pBluescript SK+. To yield SkLCS46, theSacII-XhoI insert was ligated into the corresponding SacII/XhoI restriction sites of plasmid pCAC6 (see above). The asterisks here and below indicate restriction sites generated by polymerase chain reaction. The EcoRI-XmnI fragment of Sk (nt 1007–1964) was coligated with theXmnI-AflII fragment (nt C2330-C2690) of clone CSk58 (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar), which is C (1–855), Sk (725–742), C (874–2171), into the corresponding EcoRI/AflII restriction sites (nt Sk1007/C2690) of the modified (with the cardiac II-III loop inserted)SacII-XhoI subclone of Sk (nt 86–2654) in pBluescript SK+. Finally, the SacII-XhoI insert was ligated into the corresponding SacII/XhoI restriction sites of plasmid pCAC6 (see above) to yield SkLCS18. The coding sequence of the α1 subunit of the skeletal muscle DHPR (10Tanabe T. Takeshima H. Mikami A. Flockerzi V. Takahashi H. Kangawa K. Kojima M. Matsuo H. Hirose T. Numa S. Nature. 1987; 328: 313-318Crossref PubMed Scopus (970) Google Scholar) was inserted in-frame and downstream of the coding region of a modified green fluorescence protein (GFP), cloned in a proprietary mammalian expression vector (kindly provided by P. Seeburg) as described in detail elsewhere (12Grabner M. Dirksen R.T. Beam K.G. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 1903-1908Crossref PubMed Scopus (136) Google Scholar). The SalI*-EcoRI fragment of GFP-α1S (nt 5′ polylinker-Sk1007) was coligated with theEcoRI-BglII fragments of clones SkLC, SkLCS46, and SkLCS18 (nt Sk1007-Sk4488) into the corresponding SalI*/BglII restriction sites of plasmid GFP-α1S. The Ca2+ channel β1b subunit cDNA (kindly provided by K. Campbell) was cloned as a SacI-HindIII fragment (5′ and 3′ polylinker, respectively) into the SacI/HindIII polylinker sites of the mammalian expression vector pSV-SPORT1 (LifeTechnologies, Inc.). The integrity of all the chimeric DHPRs was confirmed by sequence analysis using an ABI 377 automatic sequencer. Primary cultures of myotubes isolated from newborn dysgenic mice were prepared as described previously (13Adams B.A. Beam K.G. J. Gen. Physiol. 1989; 94: 429-444Crossref PubMed Scopus (66) Google Scholar). Approximately 1 week after plating, myotubes were microinjected (2Tanabe T. Beam K.G. Powell J.A. Numa S. Nature. 1988; 336: 134-139Crossref PubMed Scopus (584) Google Scholar) into a single nucleus with solutions of expression plasmids (300–600 ng/μl) carrying cDNAs for either GFP-α1S, GFP-SkLC, GFP-SkLCS46, or GFP-SkLCS18. Injected myotubes were subsequently examined for the development of green fluorescence. Expressing cells were evaluated for contraction (2Tanabe T. Beam K.G. Powell J.A. Numa S. Nature. 1988; 336: 134-139Crossref PubMed Scopus (584) Google Scholar) in response to electrical stimulation (80 V, 10–30 ms), macroscopic Ca2+ currents, immobilization-resistant intramembrane charge movement (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar), and subcellular channel distribution (only for GFP-SkLC). In a separate set of experiments examining the role of the β1b subunit for Ca2+ channel enhancement, dysgenic myotubes were coinjected with GFP-SkLC cDNA (600 ng/μl) and 350 ng/μl β1b-carrying mammalian expression plasmid. Additionally, dyspedic myotubes were grown in primary culture as described for dysgenic myotubes (13Adams B.A. Beam K.G. J. Gen. Physiol. 1989; 94: 429-444Crossref PubMed Scopus (66) Google Scholar) and mononuclearly injected (2Tanabe T. Beam K.G. Powell J.A. Numa S. Nature. 1988; 336: 134-139Crossref PubMed Scopus (584) Google Scholar) with 350 ng/μl β1b-carrying mammalian expression plasmid together with pure GFP vector (25 ng/μl) to enable the identification of expressing cells. Macroscopic Ca2+ currents were measured using the whole-cell patch clamp technique (15Hamill O.P. Marty A. Neher E. Sakmann B. Sigworth F.J. Pflügers Arch. Eur. J. Physiol. 1981; 391: 85-100Crossref PubMed Scopus (15174) Google Scholar). The patch pipettes (borosilicate glass) had resistances of 1.5–1.9 MΩ when filled with an internal solution containing 140 mm cesium aspartate, 10 mm Cs2-EGTA, 5 mm MgCl2, and 10 mm HEPES (pH 7.4 with CsOH). The composition of the external bath solution was 10 mm CaCl2, 145 mm tetraethylammonium chloride, 3 μmtetrodotoxin, and 10 mm HEPES (pH 7.4 with tetraethylammonium hydroxide). Test pulses were preceded by a 1-s prepulse to −30 mV to inactivate endogenous T-type Ca2+currents (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar). Test currents were corrected for linear components of leakage and capacitative currents by digitally scaling and subtracting the average of 10 preceding control currents, elicited by hyperpolarizing voltage steps (20–40 mV amplitude) applied from the holding potential of −80 mV. Ca2+ currents were normalized by linear cell capacitance (expressed in pA/pF). After the recording of whole-cell Ca2+ currents, 0.5 mmCd2+, and 0.1 mm La3+ were added to the external bath solution to enable the recording of immobilization-resistant intramembrane charge movement (gating currents). The procedure for recording and calculating maximum charge movement densities and the prepulse protocol used was described in detail elsewhere (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar, 16Adams B.A. Mori Y. Kim M.S. Tanabe T. Beam K.G. J. Gen. Physiol. 1994; 104: 985-996Crossref PubMed Scopus (17) Google Scholar). To examine the effect of Ca2+release on sarcolemmal Ca2+ current, Ca2+current and Ca2+ transients were measured (17Garcia J. Beam K.G. J. Gen. Physiol. 1994; 103: 107-123Crossref PubMed Scopus (66) Google Scholar) in normal myotubes with the external solution described above for Ca2+ currents and patch pipettes containing an internal solution composed either of 145 mm cesium glutamate, 8 mm MgATP, 0.5 mm K5-Fluo-3 (Molecular Probes, Eugene, OR), 2 mm CsCl, 10 mm EGTA, 10 mm HEPES, pH 7.4, with CsOH (10 EGTA solution) or 65 mm cesium glutamate, 5 mmMgCl2, 0.5 mm K5-Fluo-3, 40 mm BAPTA, 10 mm HEPES, pH 7.4 with CsOH (40 BAPTA solution). For the measurement of Ca2+ transients in dysgenic myotubes expressing chimeric DHPRs, the pipette contained 145 mm cesium glutamate, 8 mm MgATP, 0.5 mm K5-Fluo-3, 0.1 mm EGTA, 2 mm CsCl, 10 mm HEPES (pH 7.2 with CsOH), and the external solutions was 150 mm tetraethylammonium chloride, 10 mm HEPES, 5 mm CaCl2, 1 mm MgCl2, 1 μm tetrodotoxin (pH 7.2 with tetraethylammonium hydroxide). For the measurements of Ca2+ transients, it was not suitable to use the GFP-tagged constructs that had fluorescence excitation and emission wavelengths close to those of Fluo-3. Thus, cDNAs coding for SkLC, SkLCS46, and SkLCS18 were inserted into the expression plasmid pKCRH2 (18Mishina M. Kurosaki T. Tobimatsu T. Morimoto Y. Noda M. Yamamoto T. Terao M. Lindstrom J. Takahashi T. Kuno M. Numa S. Nature. 1984; 307: 604-608Crossref PubMed Scopus (248) Google Scholar) and were coinjected with cDNA encoding the α subunit of the human surface antigen CD8 (19Jurman M.E. Boland L.M. Liu Y. Yellen G. Biotechniques. 1994; 17: 876-880PubMed Google Scholar). Myotubes expressing the mutant channels were identified using polystyrene beads coated with CD8 antibodies as described previously (20Garcia J. Nakai J. Imoto K. Beam K.G. Biophys. J. 1997; 72: 2515-2523Abstract Full Text PDF PubMed Scopus (54) Google Scholar). Transient changes in fluorescence (ΔF) were normalized by the resting fluorescence (F). The maximum rate of change of ΔF/F was determined by fitting a line segment to the steepest portion of the transient. All recordings were made at room temperature (∼20 °C) and data are reported as mean ± S.D. GFP-SkLC-expressing dysgenic myotubes cultured on 35-mm culture dishes were superfused with a normal rodent Ringer solution (145 mm NaCl, 5 mm KCl, 2 mmCaCl2, 1 mm MgCl2, and 10 mm HEPES, pH 7.4 with NaOH) and mounted under a glass coverslip. The culture dish was subsequently fastened upside-down on the stage of a Nikon inverted microscope. Fluorescing cells were analyzed using a Sarastro 2000 confocal laser-scanning microscope (Molecular Dynamics) with a Nikon 60× PlanApo oil immersion objective (numerical aperture 1.40) and the ImageSpace™ software (Silicon Graphics Inc., Mt. View, CA). GFP excitation/emission was achieved with a filter set (488 nm/510 nm) designed for fluorescein detection. Images were 1024 × 1024 pixels with a pixel size of 0.11 μm. Step size between confocal sections was 2 μm. Images were processed using the Adobe Photoshop software (ADOBE Systems, Mountain View, CA). Previous work showed that the cardiac DHPR expressed in dysgenic (DHPR-deficient) myotubes was unable to mediate orthograde signaling (i.e. skeletal EC coupling). However, a chimera with cardiac sequence except for a skeletal II-III loop (CSk3 (5Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 344: 451-453Crossref PubMed Scopus (194) Google Scholar)) could mediate orthograde signaling. More recently, it was found that the expression of RyR-1 in dyspedic (RyR-deficient) myotubes increased the amplitude of slow L-type Ca2+ current produced by the skeletal DHPR (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar). To determine whether the II-III loop plays an important role in "receiving" this current-enhancing signal from RyR-1, we constructed a chimeric DHPR (SkLC) that was the "inverse" of CSk3: a skeletal DHPR except for a cardiac II-III loop. Electrically evoked contractions were never observed in dysgenic myotubes that had been injected with SkLC, consistent either with the possibility that the chimera was nonfunctional or that a cardiac II-III loop abolished orthograde signaling. Because electrically evoked contraction could not be used to identify myotubes expressing SkLC, we constructed a cDNA that fused GFP to the amino terminus of SkLC (Fig. 1a) to allow definitive identification by means of in situ fluorescence. Fusion proteins of this sort were shown previously not to affect the function of either muscle or brain Ca2+ channels (12Grabner M. Dirksen R.T. Beam K.G. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 1903-1908Crossref PubMed Scopus (136) Google Scholar). Myotubes expressing GFP-SkLC displayed slowly activating Ca2+currents (Fig. 2b), which were much smaller in amplitude than those present in GFP-α1S-expressing myotubes (Fig. 2a). To allow comparisons between cells, peak current-voltage relationships were fitted (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar) to yield a value of maximal Ca2+conductance (Gmax). The value ofGmax for GFP-SkLC was significantly (p < 0.005) smaller than for GFP-α1S(Table I). This decrease inGmax for GFP-SkLC did not appear to be a consequence of a reduced number of DHPRs expressed in the surface membrane because values for maximal charge movement (Qmax) were similar (p > 0.05) for GFP-SkLC and GFP-α1S (Fig. 2,d and e; Table I). The ratio of Gmax to Qmax′( Qmax′ equalsQmax minus the average, endogenous charge in dysgenic myotubes; Ref. 14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar) for GFP-SkLC was less than half that for GFP-α1S (Table I). Thus, it appears that the presence of a cardiac II-III loop prevents GFP-SkLC from receiving the current-enhancing signal from RyR-1. Indeed, the value ofGmax/ Qmax′ for GFP-SkLC was very close to the value found for dyspedic myotubes (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar), which have α1S but lack RyR-1.Figure 2Representative whole-cell Ca2+currents recorded from dysgenic myotubes expressing GFP-α1S (a), GFP-SkLC (b), and GFP-SkLCS46(c). Macroscopic Ca2+ currents were elicited by 200-ms step depolarizations from a holding potential of −80 mV to the test potentials indicated on the left (in mV). Current amplitudes were normalized by linear cell capacitance and are expressed as pA/pF. d–f, immobilization-resistant intramembrane charge movements were recorded in response to a depolarization to +40 mV following a prepulse protocol (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar). Recordings of charge movement were obtained from the same myotubes as shown above undera–c after blocking Ca2+ currents with a test solution containing 0.5 mm Cd2+ and 0.1 mm La3+. The linear cell capacitance (C) for each cell was as follows: a and d, cell b67, C = 436 pF; b and e,cell b48, C = 520 pF; c and f, cell c08, C = 588 pF.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Table ICa2+ conductance and charge movement in dyspedic myotubes and in dysgenic myotubes expressing α1S, GFP-α1S, and the GFP-SkLC clone familyData are given as mean ± SD, with the numbers in parentheses indicating the number of myotubes tested. Values ofGmax, the maximal Ca2+ conductance, were obtained by fitting the measured currents according to the functionI = Gmax(V −Vrev)/(1 + exp[−(V −VG)/kG]) (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar); I, peak current activated at test potential V;Vrev, extrapolated reversal potential;VG, potential for activation of half-maximal conductance; kG, slope factor. Values of immobilization-resistant QON were determined as described previously (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar) and were fitted according toQON = Qmax /(1 + exp[−(V −VQ)/kQ]);Qmax, maximum immobilization-resistant charge movement; V, test potential; VQ, potential at which half the charge has moved; kQ, slope factor. Q′max is the difference betweenQmax and the average, endogenous charge movementQdys(max) found in dysgenic myotubes (Qdys(max) = 2.5 nC/μF; (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar)). For all the data given, the estimated series resistance error was <10 mV. Brackets indicate two data sets compared statistically by an unpaired two-samplet test. Asterisks indicate statistically significant differences (p < 0.005), whereas no asterisk indicates p > 0.05. Values for dyspedic myotubes and for α1S-expressing dysgenic myotubes were listed for comparison and were published previously (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar, 14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar). Open table in a new tab Data are given as mean ± SD, with the numbers in parentheses indicating the number of myotubes tested. Values ofGmax, the maximal Ca2+ conductance, were obtained by fitting the measured currents according to the functionI = Gmax(V −Vrev)/(1 + exp[−(V −VG)/kG]) (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar); I, peak current activated at test potential V;Vrev, extrapolated reversal potential;VG, potential for activation of half-maximal conductance; kG, slope factor. Values of immobilization-resistant QON were determined as described previously (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar) and were fitted according toQON = Qmax /(1 + exp[−(V −VQ)/kQ]);Qmax, maximum immobilization-resistant charge movement; V, test potential; VQ, potential at which half the charge has moved; kQ, slope factor. Q′max is the difference betweenQmax and the average, endogenous charge movementQdys(max) found in dysgenic myotubes (Qdys(max) = 2.5 nC/μF; (14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar)). For all the data given, the estimated series resistance error was <10 mV. Brackets indicate two data sets compared statistically by an unpaired two-samplet test. Asterisks indicate statistically significant differences (p < 0.005), whereas no asterisk indicates p > 0.05. Values for dyspedic myotubes and for α1S-expressing dysgenic myotubes were listed for comparison and were published previously (8Nakai J. Dirksen R.T. Nguyen H.T. Pessah I.N. Beam K.G. Allen P.D. Nature. 1996; 380: 72-75Crossref PubMed Scopus (401) Google Scholar, 14Adams B.A. Tanabe T. Mikami A. Numa S. Beam K.G. Nature. 1990; 346: 569-572Crossref PubMed Scopus (222) Google Scholar). Nakai et al. (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar) previously showed that substitution of a 46-amino acid segment of skeletal sequence into the II-III loop of an otherwise cardiac DHPR produced a chimera (CSk53; α1S residues 720–765) capable of mediating strong, skeletal-type EC coupling upon expression in dysgenic myotubes. To determine whether this motif (Fig. 1 b) is also sufficient to allow reception of the Ca2+ current-enhancing signal from RyR-1, we substituted this 46-residue segment into the cardiac II-III loop of the otherwise skeletal chimera GFP-SkLC. The resulting chimera, GFP-SkLCS46 (Fig. 1 a), not only mediated skeletal-type EC coupling (electrically evoked contraction of more than half of the fluorescent cells tested in Cd2+/La3+, n > 50; data not shown) but also produced large Ca2+ current densities (Fig. 2c) with aGmax/ Qmax′ratio (>30 nS/pC) like those of GFP-α1S or α1S (Table I). Nakai et al. (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar) also showed that an otherwise cardiac chimera containing an even shorter (18-residue) skeletal segment (CSk58, α1S residues 725–742) was still able to mediate skeletal-type EC coupling; however, this coupling was weak (7Nakai J. Tanabe T. Konno T. Adams B. Beam K.G. J. Biol. Chem. 1998; 273: 24893-24896Google Scholar). To test if this 18-amino acid segment allows reception of the channel-enhancing signal from RyR-1, we constructed chimera GFP-SkLCS18 (Fig. 1, a and b). The value ofGmax for GFP-SkLCS18 was not significantly different from that of GFP-SkLC (Table I;p > 0.05). Additionally, theGmax/ Qmax′ratio for GFP-SkLCS18 was similar to that found for GFP-SkLC expressed in dysgenic myotubes or that of endogenous α1S in dyspedic myotubes that lack RyR-1 (Table I). Therefore, the minimal DHPR sequence that allows strong enhancement of Ca2+ current by RyR-1 is incomplete in, or missing from, the 18-residue skeletal segment in the II-III loop of GFP-SkLCS18. However, this minimal sequence is contained within the 46-residue skeletal segment of the GFP-SkLCS46II-III loop. The measurement ofGmax/ Qmax′provides a quantitative assessment of the strength of retrograde coupling (current enhancement from RyR-1). To obtain a similarly quantitative assessment of the strength of orthograde (EC) coupling to RyR-1, we measured depolarization-induced Ca2+ transients. Depolarization-induced Ca2+ transients were never observed with SkLC (Fig. 3 a, 0 of 13 cells tested) but were routinely observed for SkLCS46 (Fig. 3 b, 16 of 16 cells tested). The transients support the conclusion that SkLCS46 mediates skeletal-type EC coupling because they were of similar magnitude for test pulses to +30 mV (where Ca2+ current is near maximal) and +80 mV (where Ca2+ current is small as a result of reduced driving force). By the same logic, SkLCS18 was also able to mediate skeletal-type coupling because the Ca2+ transients were again similar at +30 and +80 mV (Fig. 3 c). However, the maximal rate of increase of the ΔF/F signal (at +80 mV) was only 0.023 ± 0.012 ms−1 (n = 12) for SkLCS18, which is almost 5-fold lower than the value of 0.112 ± 0.025 ms−1 (n = 16) for SkLCS46. Thus, skeletal-type coupling is much weaker for SkLCS18 than for SkLCS46. It is possible to calculate the enhancement of Ca2+ current that might have been expected for SkLCS18 relative to that measured for SkLCS46 under the assumption that there is a linear relationship between the strengths of retrograde and orthograde signaling. The enhancement of current for SkLCS46 can be defined as (G/ Q46′ − G/Q′) ÷ Q46′, where G/ Q46′ and G/Q′ are the values ofGmax/ Qmax′for GFP-SkLCS46 and GFP-SkLC, respectively. With t
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