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• W1550748049 abstract "Non-Technical Summary Calcium ions flowing through the type 1 ryanodine receptor (RyR1) calcium channel trigger contraction of skeletal muscle cells. Close to 300 mutations of the gene encoding RyR1 are responsible for several muscular diseases in human. Properties of pathological mutant RyR1s have so far been essentially assessed from studies in cultured cells and in differentiated native muscle fibres from a few available transgenic mouse models. We show that functional properties of mutant RyR1s can be reliably assessed following in vivo expression in adult mouse muscles. The Y523S, R615C and R2163H RyR1 mutants produce a similar over-sensitive activation of the calcium flux whereas I4897T RyR1 mutants are responsible for a depressed Ca2+ flux. The alterations appear to result from inherent modifications of RyR1 channel function and not from indirect changes in the muscle fibre homeostasis. The present strategy will help understand the physio-pathological defects underlying alterations of muscle function in affected patients. Abstract Mutations of the gene encoding the type 1 ryanodine receptor (RyR1) are associated with skeletal muscle disorders including malignant hyperthermia susceptibility (MHS) and central core disease (CCD). We used in vivo expression of EGFP-RyR1 constructs in fully differentiated mouse muscle fibres to characterize the function of several RyR1 mutants. Wild-type and Y523S, R615C, R2163H and I4897T mutants of RyR1 were separately expressed and found to be present within restricted regions of fibres with a pattern consistent with triadic localization. Confocal measurements of voltage-clamp-activated myoplasmic Ca2+ transients demonstrated alterations of sarcoplasmic reticulum (SR) Ca2+ release spatially correlated with the presence of exogenous RyR1s. The Y523S, R615C and R2163H RyR1 MHS-related mutants were associated with enhanced peak Ca2+ release for low and moderate levels of depolarization, whereas the I4897T CCD mutant produced a chronic reduction of peak SR Ca2+ release. For example, peak Ca2+ release in response to a depolarization to –20 mV in regions of fibres expressing Y523S and I4897T was 2.0 ± 0.3 (n= 9) and 0.46 ± 0.1 (n= 5) times the corresponding value in adjacent, non-expressing regions of the same fibre, respectively. Interestingly no significant change in the estimated total amount of Ca2+ released at the end of large depolarizing pulses was observed for any of the mutant RyR1 channels. Overall, results are consistent with an ‘inherent’ increase in RyR1 sensitivity to activation by the voltage sensor for the MHS-related RyR1 mutants and a partial failure of voltage-gated release for the CCD-related I4897T mutant, that occur with no sign of change in SR Ca2+ content. Furthermore, the results indicate that RyR1 channel density is tightly regulated even under the present conditions of forced exogenous expression. Mammalian skeletal muscle contraction is triggered by a rise in cytoplasmic Ca2+ due to Ca2+ release from the sarcoplasmic reticulum (SR) through the type 1 ryanodine receptor (RyR1). RyR1 is a massive homo-tetrameric protein, each monomer being constituted of ∼5000 residues. RyR1 channels are located in the SR membrane that faces transverse invaginations of the sarcolemma constituting the transverse (t-) tubule system. Excitation–contraction (E-C) coupling occurs within this junctional region where RyR1 gating is driven by the dihydropyridine receptor (DHPR) protein Cav1.1, which senses the changes in voltage across the t-tubule membrane (see Dulhunty, 2006). Given the primary role of RyR1 in E-C coupling, any defect of its functional properties is expected to compromise muscle force production, and thus proper motor function. Close to 300 mutations of the human RYR1 encoding gene have been identified (for review see Lyfenko et al. 2004; Treves et al. 2005; Robinson et al. 2006; Treves et al. 2008; MacLennan & Zvaritch, 2011). RYR1 gene mutations are associated in humans with malignant hyperthermia susceptibility (MHS), central core disease (CCD), multiminicore disease (MmD), centronuclear myopathy (Wilmshurst et al. 2010), atypical periodic paralysis syndromes (Zhou et al. 2010) and core-rod myopathy (Hernandez-Lain et al. 2011). Thus, there is a pressing need to better understand the mechanisms through which a given mutation alters the Ca2+ release channel function and how these defects, in turn, lead to such a diverse array of clinically distinct entities. Understanding the physiological consequences of pathological mutant forms of the RyR1 Ca2+ release channel requires the proteins to be expressed in a skeletal muscle environment where their opening and closing is controlled by the t-tubule DHPR voltage sensor. This so far has been made possible through the combined use of electrophysiology and intracellular Ca2+ measurements in the context of two main experimental strategies: expression in cultured myotubes and generation of knock-in mouse models, keeping aside (but in mind) complementary studies performed on muscle preparations from the pig model of MHS (Dietze et al. 2000; Melzer & Dietze, 2001) and from MHS patients (Struk et al. 1998). Studies of the consequences of mutant RyR1 expression in cultured RyR1-deficient (dyspedic) myotubes have led to comprehensive and widely accepted physio-pathological models of related muscle dysfunction. In brief, certain mutations render the RyR1 channel hypersensitive to activation (a hallmark of MHS) that lead to either compensated (MH in the absence of CCD) or non-compensated (MH+CCD) SR Ca2+ leak. Conversely, mutations associated with a CCD-only phenotype are proposed to reduce SR Ca2+ release independent of a change in RyR1 sensitivity or SR Ca2+ content (Avila et al. 2001, 2003; Dirksen & Avila, 2002, 2004; Lyfenko et al. 2007). In comparison with cultured myotubes, knock-in animal models offer the advantage of assessing the consequences of the mutations in fully differentiated adult muscle fibres. Mouse models are currently available for three mutant forms of RyR1: Y522S (Chelu et al. 2006), R163C (Yang et al. 2006) and I4897T (Zvaritch et al. 2007) and intracellular Ca2+ transients under voltage-clamp conditions have so far been investigated in adult muscle fibres isolated from the heterozygous Y522S and I4897T models (Andronache et al. 2009; Loy et al. 2011). Some of the results from these studies confirmed previous data from expression in myotubes, but also highlighted features specific to adult muscle fibres. In the present work we characterized the functional consequences of expressing several pathological mutant forms of RyR1 in fully differentiated normal adult mouse muscle fibres, using in vivo electroporation of cDNA constructs. As compared to the knock-in mouse models available, the present strategy was designed to bypass possible complications due to effects of the mutant channels on muscle development and maturation, to limit any masking of defects due to compensatory mechanisms, and thus, to better emphasize the primary functional deficits of the mutant channel on Ca2+ release during E-C coupling. Using a combination of voltage-clamp and confocal imaging of transfected fibres, we provide insights into the function of four different RyR1 disease mutants by directly comparing their activity with that of the endogenous wild-type RyR1s present in non-expressing regions of the same cell. All experiments and procedures were conducted in accordance with the guidelines of the local animal ethics committee of University Lyon 1, of the French Ministry of Agriculture (87/848) and of the European Community (86/609/EEC). Our experiments comply with the policies and regulations of The Journal of Physiology (Drummond, 2009) and UK regulations on animal experimentation. cDNA constructs encoding rabbit wild-type (WT) and mutant I4897T RyR1 N-terminally tagged with EGFP as well as constructs encoding the corresponding untagged R615C, Y523S and R2163H mutant of RyR1 were obtained from R. T. Dirksen's laboratory. The R615C and Y523S mutations were ligated into the WT RyR1-EGFP construct using Mlu1/Age1 restriction sites. The R2163H mutation was introduced into the WT RyR1-EGFP construct using Age1/BSiW1 restriction sites. Exogenous expression by electroporation (Trollet et al. 2006) was performed in the flexor digitorum brevis (FDB) and interosseus muscles of 4- to 6-week-old Swiss OF1 male mice using a procedure similar to the one described in detail by Di Franco et al. (2009). Mice were anaesthetized by isoflurane inhalation (3% in air, 300 ml min−1) using a commercial delivery system (Univentor 400 Anaesthesia Unit, Uninventor, Zejtun, Malta). Twenty five microlitres of a solution containing 2 mg ml-1 hyaluronidase dissolved in sterile saline was then injected into the footpads of each hind paw. Forty minutes later the mouse was re-anaesthetized by isoflurane inhalation. Plasmid DNA was then injected into the footpads of the animal at a concentration of 30 μg μl-1 in standard Tyrode solution. A 20 μl total volume of this solution was injected in different locations so as to target both the FDB and the interosseus muscles. Following the injection, two gold-plated stainless steel acupuncture needles connected to the electroporation apparatus were inserted under the skin, near the proximal and distal portion of the foot. The standard protocol used consisted in 20 pulses of 110 V cm-1 amplitude and 20 ms duration delivered at a frequency of 2 Hz by a BTX ECM 830 square wave pulse generator (Harvard Apparatus, Holliston, MA, USA). Experimental observations and measurements were carried out 2 weeks later. Single fibres were isolated from FDB and interosseus muscles using a previously described procedure (Jacquemond, 1997). In brief, mice were killed by cervical dislocation before removal of the muscles. Muscles were treated with collagenase (Sigma, type 1) for 60 min at 37°C. Single fibres were then obtained by triturating the muscles within the experimental chamber. For standard observations performed in the absence of voltage-clamp, fibres were bathed in Tyrode solution. They were dispersed on the glass bottom of either a single-well Lab-Teck chamber (Nalge Nunc, Naperville, IL, USA) or of a 50 mm wide culture μ-dish (ibidi, Munich, Germany). For intracellular Ca2+ measurements fibres were first partially insulated with silicone grease as described previously (Jacquemond, 1997). Briefly, fibres were embedded within silicone so that only a portion of the fibre extremity was left out of the silicone. Under these conditions fibres remained well maintained on the bottom of the chamber and this allowed whole-cell voltage clamp to be achieved on the silicone-free extremity of the fibre. Expressing fibres were handled with silicone so that the fibre region exhibiting EGFP fluorescence was left out of the silicone. It should be stressed that not all EGFP-positive fibres were eligible for this procedure, especially when the region of expression was within the central portion of the fibre. Once partially embedded within silicone, rhod-2 free acid was introduced into the myoplasm through local pressure micro-injection with a micropipette containing 1 mm of the dye dissolved in a solution containing 100 mm EGTA and 40 mm CaCl2 (see Solutions). Microinjection was always performed within the silicone-embedded part of the fibre, away from the silicone-free end portion under study. Following diffusion and equilibration within the cytoplasm, this was believed to achieve a final cytoplasmic concentration of rhod-2 and EGTA within the 100 μm and 10 mm range, respectively (for details concerning micro-injections see Csernoch et al. 1998). All experiments were performed at room temperature (20–22°C). The initial series of experiments were performed with a Zeiss LSM 510 laser scanning confocal microscope available at the Centre Technologique des Microstructures of University Lyon 1. The microscope was equipped with a 63× oil immersion objective (numerical aperture 1.4). Thereafter, all experiments were conducted using a Zeiss LSM 5 Exciter confocal microscope equipped with a similar 63× oil immersion objective (numerical aperture 1.4). In both cases, the EGFP excitation was provided by the 488 nm line of an argon laser and a 505 nm long pass filter was used on the detection channel. For detection of rhod-2 fluorescence, excitation was from the 543 nm line of a He–Ne laser and fluorescence was collected above 560 nm. One major aim of the experiments was to simultaneously record intracellular rhod-2 Ca2+ signals in both an EGFP-positive fibre region (enriched in exogenous RyR1s) and an adjacent EGFP-deprived region where the contribution of exogenous RyR1s should be much less. For this we specifically worked on imaging silicone-free portion of fibres locally yielding a large and restricted EGFP signal. An illustrative example is presented in Fig. 1, which shows confocal rhod-2 (top) and EGFP (middle) images from an expressing fibre with the EGFP fluorescence profile (bottom) along the yellow line superimposed to the EGFP frame. Intracellular Ca2+-related fluorescence changes were imaged by using the line-scan mode of the system with the line parallel to the longitudinal fibre axis. The majority of images were taken with a scanning frequency of 1.53 ms per line. Image processing and analysis was performed using Image/J (NIH, USA) and Origin (OriginLab Corp., Northampton, MA, USA). Confocal line-scan distribution of EGFP-tagged RyR1 channels expressed in an adult mouse muscle fibre Confocal x,y frames of rhod-2 Ca2+ indicator fluorescence (top) and from locally expressed EGFP-tagged RyR1 channels (middle) in a mouse muscle fibre. The graph depicts the fluorescence intensity distribution along the line shown in the middle panel (bottom). The scale bar on the rhod-2 image stands for the three panels. Changes in rhod-2 fluorescence were expressed as F/F0 where F0 is the resting (or baseline) fluorescence level. Changes in [Ca2+] were calculated from the rhod-2 signals using the previously described pseudo-ratio equation (Cheng et al. 1993), assuming a basal [Ca2+] of 100 nm and a Kd of rhod-2 for Ca2+ of 1.2 μm. An estimation of the Ca2+ release flux underlying the calculated global [Ca2+] transients was performed according to a previously described procedures (Collet et al. 2004; Pouvreau et al. 2006). In brief, the SR calcium release flux was calculated from the time derivative of the total myoplasmic Ca2+ obtained from the occupancy of intracellular calcium binding sites. The model included troponin C binding sites with a total concentration of sites (TNtotal) of 250 μm, an ‘on’ rate constant (kon,CaTN) of 0.0575 μm−1 ms−1 and an ‘off’ rate constant (koff,CaTN) of 0.115 ms−1; Ca–Mg binding sites on parvalbumin with a total concentration of sites (PVtotal) of 2000 μm, ‘on’ rate constant for Ca2+ (kon,CaPV) of 0.125 μm−1 ms−1, ‘off’ rate constant for Ca2+ (koff,CaPV) of 5.10−4 ms−1, ‘on’ rate constant for Mg2+ (kon,MgPV) of 3.3.10−5μm−1 ms−1, ‘off’ rate constant for Mg2+ (koff,MgPV) of 3.10−3 ms−1. Calcium transport across the SR membrane was included with a rate assumed to be proportional to the fractional occupancy of the SR pump sites with a dissociation constant (Kd,Capump) of 2 μm and a maximum pump rate of 10 μm ms−1. Resting [Mg2+] was assumed to be 1.5 mm. The model also included Ca2+-binding sites on EGTA at a concentration of 10 mm, an ‘on’ rate constant (kon,CaEGTA) of 0.056 μm−1 ms−1 and an ‘off’ rate constant (koff,CaEGTA) of 0.002 ms−1. Under the present conditions calcium binding to EGTA obviously made a predominant contribution to the calculated Ca2+ release flux as compared to intrinsic Ca2+-buffering and -removal components of the model. It should be stressed that under the present conditions where fibres were micro-injected with the dye and EGTA containing solution, it was impossible to certify that the exact same concentration of EGTA was present in all fibres. To some extent, an inherent variability in the absolute values for peak rhod-2 transients and calculated Ca2+ release flux was thus expected from fibre to fibre. For this reason we focussed our analysis on comparing the properties of Ca2+ transients and Ca2+ release within a given line-scan image, between regions of the same fibre with and without detectable EGFP-RyR1s channels. Indeed, throughout an entire given fibre identical concentrations of EGTA as well as other intrinsic mobile buffers were expected. We also implicitly assumed that the level of intrinsic immobile Ca2+-binding proteins, pumps and channel proteins involved in Ca2+ homeostasis were spatially homogeneous throughout the fibre. An RK-400 patch-clamp amplifier (Bio-Logic, Claix, France) was used in whole-cell voltage-clamp configuration. Fibres were bathed in a TEA-containing extracellular solution (see Solutions). In the series of experiments performed with the Zeiss LSM 510 microscope, command voltage pulse generation was achieved with an SMP300 voltage pulse generator (Bio-Logic). For the experiments with the Zeiss LSM 5 Exciter microscope, an analog–digital converter (Digidata 1440A, Axon Instruments, Union City, CA, USA) controlled by pCLAMP 9 software (Axon Instruments) was used. Voltage-clamp was performed with a microelectrode filled with a solution that mimics ionic conditions of the intracellular environment (see Solutions). The tip of the microelectrode was inserted through the silicone, within the insulated part of the fibre. Analog compensation was systematically used to decrease the effective series resistance. Membrane depolarizing steps of 0.5 s duration were applied from a holding command potential of –80 mV. FDB and interosseus muscles were electroporated with the EGFP-I4897T-RyR1 construct. Two weeks later, single fibres were isolated using collagenase treatment as described above. Muscles were triturated in Tyrode solution onto glass slides. The presence of EGFP-positive fibres on the slides was checked by fluorescence microscopy. Slides were air-dried and muscle fibres were fixed in cold methanol (–20°C) for 10 min. Slides were then stored at –80°C. Slides were re-hydrated in phosphate-buffered saline (PBS), blocked for 50 min at room temperature (M.O.M. blocking kit, Vector Laboratories) and incubated overnight with a mouse anti-RyR monoclonal antibody (34C). Slides were washed for 5 min in PBS, incubated for 1 h with an anti-mouse Cy3-conjugated secondary antibody and then washed 6 times in PBS before being mounted with a coverslip. Observations were made with a Zeiss LSM 5 Exciter confocal microscope using a 63× oil immersion objective (numerical aperture 1.4). The EGFP excitation was provided by the 488 nm line of an argon laser and a 505–530 nm band-pass filter was used on the detection channel, while Cy3 fluorescence was excited with the 543 nm line of a He–Ne laser and fluorescence was collected above 560 nm. The intracellular pipette solution contained (in mm) 120 potassium glutamate, 5 Na2-ATP, 5 Na2-phosphocreatine, 5.5 MgCl2, 5 glucose, 5 Hepes. The injection solution contained (in mm) 100 EGTA, 40 CaCl2 and 1 rhod-2 (tripotassium salt). The extracellular solution used in whole-cell voltage-clamp recordings contained (in mm) 140 TEA-methanesulfonate, 2.5 CaCl2, 2 MgCl2, 10 TEA-Hepes and 0.002 tetrodotoxin. All solutions were adjusted to pH 7.20. Least-squares fits were performed using a Marquardt–Levenberg algorithm routine included in Origin. Data values are presented as means ± SEM for n fibres. Statistical significance was determined using Student's t test assuming significance for P < 0.05. EGFP fluorescence images revealed that all exogenous RyR1s tested in the present study exhibited similar properties in terms of subcellular localization when expressed in adult skeletal muscle fibres: they were present locally within one or a few spatially restricted regions, near a nucleus, as previously reported and illustrated for RyR3 channels under similar expression conditions (see Fig. 1 in Legrand et al. 2008). In terms of transverse distribution, the GFP-tagged channels were found to occupy a very substantial volume of the fibre interior, commonly covering a distance of several tens of micrometres inside the fibre. The yield of expressing fibres was always low with the number of EGFP-RyR1 positive fibres rarely exceeding 20–30 in a given FDB or interosseus muscle. Although no systematic quantification was made, this number did not appear to change depending on the construct; in other words none of the RyR1 mutants seemed to be expressed in a specifically more or less frequent manner. Figure 2 presents illustrative confocal (x,y) images of rhod-2 (left) and EGFP (right) fluorescence collected from separate muscle fibres expressing (from top to bottom) the WT-RyR1, Y523S-RyR1, R615C-RyR1, R2163H-RyR1 and I4897T-RyR1 construct, respectively. The pair of graphs for each construct depicts average EGFP (top) and rhod-2 (bottom) fluorescence profiles along the horizontal axis of the area highlighted by a box in each EGFP image. Fluorescence profiles are shown normalized to the maximum value. For each construct, EGFP yields a succession of double rows separated by a region of lower fluorescence, the distance between two consecutive double-row peaks being ∼2 μm. This distribution is consistent with physiological positioning of the expressed RyR1 channels at the triad and indicates that any observed alteration in SR Ca2+ release was unlikely to be related to severe RyR1 channel mis-targeting. The rhod-2 fluorescence also exhibited a distinct repetitive striated pattern throughout the entire fibres, irrespective of the presence of expressed RyR1s; rhod-2 fluorescence was maximal near the centre of the EGFP double peaks, corresponding to the position of the Z-line. This is likely to result from rhod-2 binding to structural sarcomeric proteins but also, to some extent, from rhod-2 accumulation into mitochondria. Possible related complications regarding the interpretation of the voltage-activated fluorescence transients were not considered. Rhod-2 was also readily detected within nuclei where it appeared to distinctively stain nucleoli, as illustrated in the image from the fibre expressing wild-type RyR1 channels. Pattern of EGFP-tagged WT and mutant RyR1 channel expression in mouse muscle fibres Representative examples of the expression pattern of (from top to bottom) WT-RyR1, Y523S-RyR1, R615C-RyR1, R2163H-RyR1 and I4897T-RyR1. For each example, the images correspond to the rhod-2 and EGFP fluorescence in the expressing fibre. The top and bottom traces in the graphs on the right show the normalized fluorescence intensity profiles within the boxed region shown in the corresponding EGFP image. One critical issue of the present experimental strategy was whether exogenous RyR1 expression per se would alter the functional properties of SR Ca2+ release. This could be thought to occur if, for instance, an unphysiologically large population of RyR1s were present and active in the SR membrane of the transfected fibres. Therefore we quantified the effect of expressing wild-type RyR1 channels under the present conditions. Figure 3A shows rhod-2 line-scan images recorded in a fibre expressing EGFP-WT-RyR1 channels; the fibre was depolarized by 0.5 s-long steps from –80 mV to the indicated potentials. The EGFP fluorescence profile along the scanned line is shown on the right side of each image; the black and the red double-arrows near the top profile indicate the x position where the rhod-2 fluorescence signals were measured in order to generate the traces shown in Fig. 3B. The rhod-2 signal was averaged over 50 adjacent rows centred at these two positions. The left panel in Fig. 3B shows the corresponding rhod-2 F/F0 signals: the red F/F0 traces were from the region of highest expression (red arrow) of EGFP-WT-RyR1 channels whereas the black traces were from the region of lowest expression (black arrow). The right panel in Fig. 3B shows the corresponding Ca2+ release flux traces calculated as described in Methods. The flux exhibits a peak followed by a more or less complex decay phase leading to a low steady level, in agreement with recent observations in mouse FDB muscle fibres stimulated by long-lasting voltage-clamp depolarizations (Royer et al. 2010). Figure 3B shows that traces recorded from the high-expression line region (red) yielded a time course very similar to the one of the low-expression region but their amplitude was slightly depressed. The voltage dependences of peak F/F0 and peak Ca2+ release flux from that fibre are shown in Fig. 3C and D while Fig. 3E shows the voltage dependence of the estimated total released Ca2+ (expressed in terms of myoplamic volume) calculated from the time integral of the Ca2+ release flux traces. Values for this parameter also tended to be slightly depressed in the region of highest expression of exogenous wild-type RyR1s. Voltage-activated SR Ca2+ release in EGFP-tagged WT-RyR1-expressing fibres A, confocal (x,t) line-scan images of rhod-2 fluorescence taken from a region of a muscle fibre that included an area of high-expression of EGFP-tagged WT-RyR1 channels. The fibre was depolarized by 0.5 s-long test pulses from –80 mV to the indicated potentials. The graph on the right of each image shows the EGFP fluorescence intensity profile along the line. The red and the black double-arrows indicate the region of the line from which the red and black traces shown in B were calculated. B, time course of rhod-2 F/F0 fluorescence (left) and of the corresponding calculated Ca2+ release flux (right) for the two regions indicated by double-arrows in A. The thick black and thin red traces are from the low-expression and high-expression regions, respectively. C, voltage dependence of the initial peak F/F0 rhod-2 fluorescence in the high-expression (open circles) and low-expression (filled circles) regions of the same fibre. D and E corresponding voltage dependence of the peak Ca2+ release flux and total released Ca2+, respectively. F and G comparison of peak Ca2+ release flux and of the total released Ca2+ in regions of a muscle fibre exhibiting high and low WT-RyR1 expression. Values for the two parameters were measured from traces as shown in Fig. 3B. F, voltage dependence of the average (±SEM) ratio of peak Ca2+ release measured in the high-expression region to the corresponding value in the low-expression region of a same line-scan image. G, voltage dependence of the average (±SEM) ratio of total released Ca2+ in the high-expression region to the corresponding value in the low-expression region of a same line-scan image. For depolarization to potentials from –30 to +10 mV, number of fibres is 7, 11, 10, 10 and 8, respectively. Fibres were from 5 distinct mice. Statistical significance indicates that the mean ratio value is significantly different from 1 (*P < 0.05). It should be stressed that throughout the present work, one reproducible feature of the Ca2+ release flux time course was that the decay phase tended to be globally faster as the pulse amplitude was increased. This resulted in the voltage dependence of total released Ca2+ tending to reach a plateau level for lower levels of membrane depolarization than the peak Ca2+ release flux. Although changes in SR Ca2+ permeability and/or SR buffering power may influence the exact time course of the Ca2+ release flux (and thus the total amount of released Ca2+) during these long pulses (see for instance Royer et al. 2010), the late phase of SR Ca2+ release decay is believed to depend largely upon SR Ca2+ depletion. Thus, SR Ca2+ content should have a strong influence on the estimated total released Ca2+. As a simplifying approximation, we assumed that any severe change in the SR Ca2+ content due to the specific conditions tested here would be reflected as an alteration of the plateau value of total released Ca2+ measured for large levels of depolarization. As discussed in Methods, a comparison of mean values for Ca2+ transient parameters between distinct batches of fibres was complicated by potential fibre to fibre differences in intracellular EGTA concentration. Thus, we essentially restricted the statistical analysis of effects of exogenous RyR1 channels to the value of the ratio between peak Ca2+ release flux in EGFP-rich and EGFP-poor regions of a same line-scan image and to the corresponding ratio value for total released Ca2+. The ratio should be equal to unity if the value for peak Ca2+ release (or total released Ca2+) is identical in the two different regions of the line-scan. Statistical difference was tested versus the hypothesis of the ratio being 1. These ratio values were only evaluated when there was a clear Ca2+ transient in the EGFP-rich and EGFP-poor regions of the scanned line. Figure 3F shows the voltage dependence of the mean (±SEM) ratio for peak Ca2+ release flux in the region of highest expression of wild-type RyR1s relative to the corresponding value in the region of lowest RyR1 expression along the same line. Figure 3G shows the corresponding voltage dependence of the mean ratio for estimated total released Ca2+. For voltages more positive than –20 mV, peak Ca2+ release tended to be somewhat smaller in regions yielding the largest EGFP signal, though this was statistically significant only at +10 mV. Conversely, ratio values for total released Ca2+ were significantly smaller than unity (by ∼10–15%) for all potentials greater than –30 mV. Overall, results in Fig. 3 indicate that expression of wild-type EGFP-RyR1 channels was not entirely without consequence on voltage-activated SR Ca2+ release: a tendency for a modest reduction in peak rate and total amount of released Ca2+ was observed in fibre regions yielding the highe" @default.
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• W1550748049 date "2011-11-01" @default.
• W1550748049 modified "2023-09-24" @default.
• W1550748049 title "Defects in Ca²⁺release associated with local expression of pathological ryanodine receptors in mouse muscle fibres" @default.
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• W1550748049 doi "https://doi.org/10.1113/jphysiol.2011.216408" @default.
• W1550748049 hasPubMedCentralId "https://www.ncbi.nlm.nih.gov/pmc/articles/3240878" @default.
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