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MUSCLE CELL BIOLOGY AND CELL MOTILITY
1Section of Cellular Signaling, Department of Molecular Biophysics and Physiology, Rush University, Chicago; and 2Department of Physiology and Biophysics, University of Illinois at Chicago, Chicago, Illinois
Submitted 3 December 2004 ; accepted in final form 30 August 2005
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ABSTRACT |
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 TOP ABSTRACT METHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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excitation-contraction coupling; sarcoplasmic reticulum; ryanodine receptors; Ca2+ imaging
Initial indications of a basal inhibition by DHPRs originated from comparisons of excitation-contraction (EC) coupling between amphibian and mammalian muscle. The comparisons led Shirokova et al. (51) and Zhou et al. (67) to conclude, in agreement with earlier studies (15, 16), that CICR has a limited or null role in mammalian muscle. An important contribution of CICR, however, was upheld for frog muscle.
The apparent differences in the contributions of CICR between amphibians and mammals ought to be explained by the different molecular makeup and supramolecular assembly of their EC coupling devices. Whereas frog fast-twitch muscles contain the 
- and 
-isoforms, in approximately equal densities (42, 56), most muscles of adult mammals exclusively express the -homolog, RyR1 (19, 34, 58), whereas some additionally express the -homolog, RyR3, in minor amounts.
According to Felder and Franzini-Armstrong (17), RyRs form separate arrays of channels, placed parajunctionally, on the sides of the triadic junction. Junctional clusters are instead constituted exclusively by the (or, in mammals, the RyR1) isoform. Because only the latter are associated with tetrads of DHPRs in the T tubule membrane (3), the activation mechanisms of these two sets of channels must be distinct (67). The parajunctional channels, which lack directly interacting DHPRs, can be under voltage control only indirectly, presumably by CICR. Conversely, the location of junctional channels should allow them to be directly controlled by voltage sensors. In agreement with these functional expectations, studies in 1B5 RyR-deficient myotubes show that RyR1 can be controlled by membrane voltage, whereas RyR3, if expressed alone, can only be activated via CICR (18).
The notion of a basal inhibition originated in good measure from the very different incidence of Ca2+ sparks in muscles of frogs and mammals. Ca2+ sparks are discrete local elevations of Ca2+ concentration ([Ca2+]), which reflect opening of a group of RyR channels spontaneously or in response to voltage or other stimuli (6, 61). The activation of multiple channels in Ca2+ sparks is believed to involve Ca2+ as mediator. This consensus derives largely from evidence collected in frog muscle, including that sparks are promoted by increases in resting cytosolic [Ca2+] ([Ca2+]cyto) (28), by increases in SR Ca2+ ([Ca2+]SR), and by caffeine (24), which increases RyR sensitivity to activation by Ca2+ (15, 35).
In contrast with amphibians, sparks are rarely observed (10) or not found at all (66) in intact adult skeletal muscle cells of mammals, and are not elicited in adult cells by voltage-clamp depolarization (11, 52). Spark frequency, however, becomes much greater under other conditions, including membrane permeabilization by saponin or its removal by "peeling" (27, 66) and interference with mitochondrial function (26). Therefore, sparks are possible in the mammal, but appear to be suppressed. Comparisons of spark frequency in 1B5 dyspedic cells expressing alternatively either isoform demonstrated that the suppression affects largely isoform 1 (62).
Suppression of a related sort was revealed by Murayama and Ogawa (37, 38) affecting bovine RyR1 and amphibian -isoforms. Because was not affected, the stabilization may explain the greater contribution of CICR in the amphibian. The authors provided evidence pointing at FKBP12 as a possible effector of this inhibition in mammalian muscle.
The inhibition has an intriguing functional feature. Using primary mammalian myotubes, Shirokova et al. (53) showed dual spatially segregated manifestations of Ca2+ release. Whereas some regions of the myotube featured strict control of Ca2+ release by membrane voltage (of the "skeletal" type, i.e., not dependent on Ca2+ entry), other regions did not respond to voltage, but produced Ca2+ sparks essentially unrelated to the depolarizing pulses. These two forms of Ca2+ release appear to be mutually exclusive (regions that respond to voltage do not produce sparks), which suggests that something in the devices underpinning voltage control, perhaps the DHPR itself, suppresses spark generation. Consistent with these ideas, Lee et al. (33) provided evidence of an inhibitory allosteric effect in 1B5 myotubes. A putative molecular locus underpinning the suppression is indicated by the existence of a malignant hyperthermia-inducing mutation in the III-IV loop of the DHPR in humans (65).
Motivated by these varied observations, the present study set out to test whether T tubules play a role in the suppression of sparks. We also explored, using myotubes lacking DHPRs, whether the voltage sensors themselves are needed for the inhibition. Because the production of sparks in mdg muscle required the use of special nonphysiological conditions, we used the shifted excitation and emission ratio (SEER) imaging technique (31) to compare [Ca2+]cyto and intrastore [Ca2+] in both types of cells under those conditions.
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METHODS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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Muscle tissue was dissected from the limbs and digested with 2 mg/ml collagenase I (Worthington Biochemical, Lakewood, NJ) in 0 Ca2+-0 Mg2+ PBS (GIBCO, Carlsbad, CA) at 37°C for 30 min. Myoblasts were collected by passing the suspension through a 40-µm-thick nylon filter (Falcon, Bedford, MA), seeded in glass-bottomed microwell dishes (MatTek, Ashland, MA), and maintained in the growth medium (DMEM, 20% FBS, GIBCO) for 2 days before they differentiated into myotubes in the differentiation medium (DMEM, 2.5% horse serum). Images were obtained from cultures that underwent differentiation for 2–4 days. Experiments were performed in compliance with National Institutes of Health guidelines and were approved by the Institutional Animal Care and Use Committee of Rush University.
Solutions. The Krebs solution was composed of (in mM) 136 NaCl, 5 KCl, 2.5 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.0. High-Ca2+ Krebs (K/Ca) was composed of (in mM) 118 NaCl, 5 KCl, 26 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.0, caffeine (K/Ca+C), with 50 µM nifedipine (K/Ca+N), or both (K/Ca+NC), were added for different purposes. All solutions contained 1 µM tetrodotoxin to prevent spontaneous activation.
For dye loading, cell cultures were immersed in Krebs solution with 5 µM of either indo-1 AM or mag-indo-1 AM. For membrane permeabilization, cultures were immersed in an internal solution containing (in mM) 130 K-glutamate, 10 Trizma maleate, 5 Na2ATP, 10 Tris-PC, 1 EGTA, 5 glucose, 8% dextran, 0.1 CaCl2, 7.23 MgCl2 (free Ca2+ = 50 nM, free Mg2+ = 1 mM), and 0.05 rhod-2 plus 0.002% saponin.
The solutions were prepared and used at 18–20°C. Chemicals for these solutions were purchased from Sigma-Aldrich (St. Louis, MO).
Optical measurements and data analysis. For detection of spontaneous Ca2+ sparks, cultures were incubated with 5 µM fluo-4 AM (Molecular Probes, Eugene, OR) in Krebs solution for 50 min at 20°C, then washed and maintained in fresh Krebs solution for an additional 40 min. For double-staining experiments, the cultures were exposed to 10 µM 4-{2-[6-(dioctylamino)-2-naphthalenyl]ethenyl}1-(3-sulfopropyl)-pyridinium (di-8-ANEPPS) (Molecular Probes) for <15 min to minimize its entry into organelles. The dye was then washed away with Krebs solution.
Cells were imaged with a confocal scanner equipped with a x40 water-immersion objective (model MRC-1000, Zeiss; numerical aperture 1.2). Dual images were obtained with excitation light of 488 nm and simultaneous recording at 
= 530 nm (±25-nm bandwidth) for detection of cytosolic Ca2+ events, and >588 nm for detection of structures stained by di-8-ANEPPS. Ca2+ events were detected automatically in xt or xy images and normalized as described by Brum et al. (4). The detection program carries out automatic determination of spark parameters: amplitude (peak F/F0), full width above half magnitude (FWHM), full duration above half magnitude (FDHM), and rise time (from 10% to full peak at the spatial center of the spark) (66).
The location of sparks in myotubes relative to the surface membrane was represented by a number that quantified it relative to the cell radius, with 0 corresponding to the membrane and 1 to the cell center. The fractional radius of the myotube invaded by T tubules (fT) was determined by comparison of T-tubule-free and occupied areas (dashed line in Fig. 1B) with midcell xy images of doubly stained myotubes.
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SEER (31) required the simultaneous acquisition of two confocal fluorescence images (named F11 and F22) produced by alternating line by line ("line interleaving") two excitation lights (351 and 364 nm) and two fluorescence emission ranges (390–440 nm and 465–535 nm). A third image, F33, was triply interleaved when rhod-2 was present in the perfusion medium, for the purpose of demonstrating permeabilization of the plasmalemma. F33 was excited at 543 nm and acquired at 550–630 nm. Imaging was done with the confocal system (model TCS SP2; Leica Microsystems, Exton, PA), using a x63 water-immersion objective, numerical aperture 1.2. Examples of images F11(x,y) and F22(x,y) of cells loaded with indo-1 AM are provided in Fig. 6, A and B. Images of a permeabilized cell loaded with mag-indo-1AM are shown in Figs. 9, A and B, and 10B. F33 is shown in Figs. 9 and 10. All other images are of the xy ratio [R(x,y)], defined as F11(x,y)/F22(x,y), shown in areas considered to be well stained (in other areas color was set to gray). The well-stained areas were those with dye concentration comprised between the mean and the mean plus three times the standard deviation of the pixel-by-pixel distribution of dye concentration within the image. Dye concentration was calculated from a linear combination of F11 and F22 (31).
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 | (1) |
is the ratio of F22 at [Ca2+] = 0 to F22 at saturating [Ca2+]. Parameter values are given by Launikonis et al. (31). Equation 1 and the equation published by Grynkiewicz et al. (25) differ only in the definition of .
Measurement of [Ca2+]cyto.
For the measurement of [Ca2+]cyto with indo-1, the culture dishes were loaded and then imaged intact. As with any procedure on intact cells loaded with a dye in AM form, there will be some interference by the dye that accumulates inside organelles. A correction for this error, developed in the APPENDIX, leads to an equation isomorphic with 1
 | (2) |
Calibration experiments to determine the parameters of Eq. 2 were carried out in cultures loaded with indo-1, so that the organelles would cause similar interference as in the intact cells. After incubation with indo-1 AM, the loaded cells were membrane permeabilized and SEER imaged in solutions with known buffered [Ca2+] and 50 µM indo-1. Keff was determined by fitting Eq. 2 to the data points (R, [Ca2+]), where R was the ratio averaged in those areas that satisfy a criterion for good staining described above. The parameter values were R0 = 0.51, Rmax = 3.94, Keff = 1,620 nM, and = 4.11, implying that Keff was 394 nM.
Statistics. Line graphs, such as Fig. 6F, plot averages of SEER ratio values over all well-stained pixels of a cell or cells within an image. The standard errors of such means are uniformly smaller than the symbol size. When a bar is plotted, the symbols represent means over averages of pieces of a cell or image. In those cases, the bar covers means ± 2 SE. Such piecewise approach was necessary to exclude well-stained debris, the inclusion of which would have biased the average. Significance of differences indicated in vertical bar graphs is determined by two-tailed t-tests.
Immunofluorescence staining. Cultures were fixed for 20 min at –20°C with methanol-acetone (1:1) precooled at –20°C. Fixed cells were incubated with blocking solution overnight at 4°C or 1 h at room temperature. The blocking solution was PBS (GIBCO) containing 10% normal goat serum, 1% BSA, and 0.1% Triton X-100 (Sigma-Aldrich). The cells were incubated with the primary antibodies, anti-RyR1 (1:1,000) or anti-RyR3 (1:2,000), diluted in the blocking solution for 1 h at room temperature or overnight at 4°C, then incubated with secondary antibodies, Cy2-conjugated goat anti-rabbit (Jackson ImmunoResearch Laboratories, West Grove, PA), in the blocking solution for 45 min at room temperature. Control experiments were done by replacing primary antibody with 1:1,000 normal rabbit serum (Sigma-Aldrich) in blocking solution. The primary antibodies, a gift from Dr. V. Sorrentino (University of Siena, Italy), have been shown not to cross-react with each other (22).
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RESULTS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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In myotubes produced in primary cultures from normal newborn mice, Ca2+ sparks were observed consistently at a low frequency. A total of 79 sparks were detected and characterized in xy images from 17 intact cells immersed in Krebs solution. Sparks occurred more frequently in myotubes of larger diameter, such as the one shown in Fig. 1. The culture was dually stained with fluo-4 AM and the membrane-soluble dye di-8-ANEPPS, and images were acquired simultaneously at two emission bands. In Fig. 1A, the fluorescence of fluo-4 in one image is normalized to the average of six other images from the same region of the specimen. Figure 1B is the averaged fluorescence of di-8-ANEPPS at longer wavelengths in the same region. It allows visualization of T tubules.
Like the one shown in Fig. 1, most Ca2+ sparks occurred away from the peripheral region. As shown in Fig. 1B, regions where sparks were frequent did not have structures stained with the membrane marker. A numerical location of the events recorded in these images was defined as the ratio (distance from center of spark to cell edge)/(cell radius). Figure 2 shows the histogram of locations of all identified sparks.
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Topographic pattern of spark generation is lost in dysgenic myotubes. If, as shown, the presence of T tubules suppresses the generation of Ca2+ sparks, a first candidate as molecular inhibitor ought to be the DHPRs, which interact mechanically with RyR1 channels in the T-SR junction (40). Skeletal muscle cells from mice homozygous for the mdg mutation (47), which lack DHPRs in the T tubules (29), were used to test this hypothesis. Primary cultures of skeletal muscle taken from neonatal mdg+/+ pups produced myotubes that were morphologically similar to those of the wild type, but appeared to have a somewhat greater diameter, as described by Powell et al. (48). Figure 3A shows the image of an mdg cell labeled with di-8-ANEPPS. After 3–4 days in differentiation medium, dysgenic myotubes, like their wild-type counterparts, had a partially developed T tubule structure, occupying the peripheral region to an extent similar to that in normal myotubes (fT = 0.234 ± 0.03, n = 8).
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To induce spark production, the cultures were exposed to K/Ca+C, a modified Krebs with 26 mM Ca2+ and a low dose of caffeine (1 mM). As shown in Fig. 3B, 315 sparks were detected and characterized in xy images from 24 dysgenic myotubes. The most remarkable difference with the normal cells was that sparks could occur just inside the plasmalemma.
As shown in Fig. 3A, T tubules could be visualized well in cells stained with di-8-ANEPPS. The spatial location of the events, represented by the histogram in Fig. 4, bore little or no relation with the location of T tubules (represented by the horizontal bar). Therefore, in dysgenic cells, the suppression effect on the releasing channel opening was lost under the conditions required to observe sparks. This absence could reflect a fundamental molecular influence or just a trivial artifact.
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The histogram of locations of 361 sparks from 26 normal myotubes in K/Ca+NC is plotted in Fig. 5B. It shows that the pattern of avoidance of T tubules by sparks, which was lost in mdg cells, persisted in normal cells under these conditions. The averaged morphological parameters of 165 events detected in xt images (see Table 1). The similarity of spark amplitudes in mdg and wild type suggests that a rough parity of cytosolic and intra-SR Ca2+ prevailed under these conditions, which proved to be true in direct measurements described next. The result is consistent with the interpretation that DHPRs are required to preserve the spatial segregation of T tubules and Ca2+ sparks.
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Evolution of [Ca2+]cyto. Figure 6, A–C, shows images F11, F22, and R (=F11/F22) of wild-type myotubes in Krebs solution. Figure 6, D and E, show ratio images in subsequent stages of work with the same culture dish. The average ratio in Fig. 6C corresponds to a [Ca2+]cyto of 152 nM. When the solution was changed to K/Ca with 50 µM nifedipine and 1 mM caffeine, the ratio did not change (Fig. 6D). After a later change to K/Ca without nifedipine, the ratio increased substantially, eventually reaching a value corresponding to [Ca2+]cyto of 1.3 µM (Fig. 6E). As shown, [Ca2+]cyto could then be very different in different cells. Figure 6F plots the time course of image-averaged ratio (the right-side axis is labeled with the corresponding [Ca2+] values, calculated using Eq. 2 in METHODS).
Corresponding ratio images for cultures of dysgenic cells are illustrated in Fig. 7. [Ca2+]cyto in Krebs varied near 170 nM. Upon changing the solution to 26 mM (no caffeine), [Ca2+]cyto remained close to initial values. The addition of 1 mM caffeine appeared to cause a small increase in some cells. Figure 7D plots the time course of image-averaged ratio and [Ca2+]cyto.
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Comparisons of SR luminal [Ca2+]. Figure 9 illustrates the technique. Figure 9, A and B, are images of a cell in a culture dish loaded with mag-indo-1 AM and then membrane-permeabilized. Figure 9C is the ratio F11/F22, restricted as described in METHODS to areas of the cell where the dye concentration was above the cellular mean. This restriction essentially removes nuclei, as well as debris, from the ratio image. The image in Fig. 9D is of fluorescence F33 of the dye rhod-2, present in the solution to identify permeabilized myotubes.
In principle, the permeabilization step reduces one's ability to follow changes in [Ca2+]SR upon changing extracellular solutions because the SR contents may vary after the plasma membrane is permeabilized. The approach illustrated in Fig. 10 was designed to evaluate the evolution of [Ca2+]SR after permeabilization. Figure 10A shows fluorescence of rhod-2, in an mdg culture dish preloaded with mag-indo-1 AM and immersed in the permeabilizing solution (which contains 50 nM free Ca2+). Figure 10B is fluorescence F22 of mag-indo-1, which is present in cytosol and organelles. A very bright cell in Fig. 10B corresponds to a dark region of Fig. 10A (dashed contour), marking a myotube that has not been permeabilized. The myotube that is highly fluorescent in Fig. 10A has much lower F22 intensity in Fig. 10B. The cellwide concentration of mag-indo drops 
10 times after permeabilization, consistent with the presence of much more dye in the cytosol than in organelles. Figure 10C shows the ratio with a high value in the permeabilized cell, consistent with the high [Ca2+] in the SR, and a low value in the intact cell, where the dye signal is largely determined by [Ca2+]cyto.
The heterogeneous staining pattern of Fig. 10, A–C, is characteristic of initial stages of permeabilization. Although it is not possible to know the exact time, most cells become permeabilized in 2–3 min. The evolution of [Ca2+]SR can then be followed for tens of minutes after permeabilization, as illustrated in Fig. 10, D–F. Figure 10D is from a wild-type culture in Krebs, and Fig. 10E is from a different wild-type dish, exposed to K/Ca for 10 min before permeabilization. The evolution of [Ca2+]SR after permeabilization, plotted in Fig. 10F, is slow. Hence, there is time after permeabilization to obtain several images and derive an average R that will represent approximately the prevailing level of [Ca2+]SR in the conditions imposed before permeabilization. The plot in red is representative of the substantial increase in [Ca2+]SR observed in normal myotubes in high Ca2+ Krebs.
Figure 11 summarizes the measurements of [Ca2+]SR. The ratio measured in wild-type cells immersed in Krebs corresponds to a [Ca2+]SR of 82 µM. Immersion in K+/Ca2+ with nifedipine increased [Ca2+]SR significantly, to 144 µM. As expected, immersion in high [Ca2+]o without the dihydropyridine caused an even greater increase in [Ca2+]SR, to 284 µM. The [Ca2+]SR level of mdg cells in Krebs, 206 µM, was significantly greater than that of normal cells in the same condition. Immersion of mdg cultures in K+/Ca2+ had an unexpected result, [Ca2+]SR decreased. Serendipitously, this decrease made the average value in mdg cells identical to that of wild-type cells in K/Ca+N (K/Ca and K/Ca+N were the conditions used for comparing location of sparks in mdg and wild-type cells).
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DISCUSSION |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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20% of the radius, where T-tubular structures are present and Ca2+ sparks are largely absent. The study also found evidence that DHPRs in the T tubules appear to have a crucial role in this inhibitory effect. T tubule structures are essential for preventing Ca2+ sparks. Ca2+ sparks of skeletal muscle result from the brief concerted opening of clusters of release channels (23). In mammalian muscle, few Ca2+ sparks were observed in fibers with intact plasmalemma (11). To induce Ca2+ sparks in mammalian muscle, the T tubule membrane had to be permeabilized by application of saponin (27, 66). This suggests that the integrity of the T tubule structure may be necessary to prevent spontaneous opening of release channels. Embryonic myotubes, which express Ca2+ release channels but have an incompletely developed T tubular structure, were used to test this hypothesis. With the use of a double-staining technique, Ca2+ sparks and T tubule structures were imaged simultaneously. First, it was demonstrated that the regions that gave rise to sparks were essentially devoid of T tubules. Conversely, the regions that did not produce sparks correspond to areas of the cell that had T tubules. It was also shown that the absence of sparks in the T tubular regions was not due to a lack of expression of RyR isoforms, of either sort.
The present work shows that T tubules and sparks occupy mutually exclusive spaces. As a consequence, the suppression of sparks must be due to T tubule-associated components, or to structures that are modified when the T tubule integrity is affected. At the molecular level, the suppression must be due to an interaction between release channels and junctional structures of some sort. The conclusion is consistent with the rapid decrease in frequency of spontaneous Ca2+ sparks in mouse muscle observed in the first two postnatal weeks (8), at a time when development of T tubules is becoming complete (57). It is also consistent with the existence of a malignant hyperthermia-inducing mutation in the III-IV loop of the DHPR in humans (65), which presumably weakens a basal inhibitory effect on RyR channels.
No T-SR docking is required for production of Ca2+ sparks. According to Takekura et al. (57), ontogenesis of Ca2+ release units requires docking of SR to embryonic plasmalemma or T tubules, to permit local clustering of release channels and voltage sensors. In contrast with this view, the present results demonstrate the presence of functional sources of Ca2+, capable of generating sparks in regions of SR not yet docked to the T tubule network. It follows that clusters of channels capable of activation form at stages before T-SR docking. Therefore, docking does not appear to be necessary for development of a spark-capable cluster. Instead, the effect of linkage with T tubules appears to be to put such clusters under a strong inhibitory control, which suppresses their ability to fire under normal conditions.
While these "unmoored" regions of SR generate sparks, the events are profoundly different from sparks observed in adult muscle (67). Most notably, their spatial width is nearly threefold greater in myotubes, which is only possible if the activation propagates over large, extensive groups of channels. Again, this is an indication of a susceptibility to activation that is largely lost on docking with T tubules.
DHPRs are necessary for basal suppression of Ca2+ sparks.
DHPRs in the T tubule membrane function as activators, which open release channels in the SR membrane during an action potential through a direct interaction. Some studies have also shown that DHPRs may function as inhibitors for the Ca2+ release channels, including the work of Suda (55), indicating possible involvement of DHPRs in terminating Ca2+ release in rat skeletal myotubes, blockage of Ca2+ release from SR vesicles by a peptide derived from the II-III loop of the DHPR (13), an effect of DHPR channel blockers on caffeine sensitivity of 1B5 myotubes expressing RyR1 (33) and the demonstration that the R1086H mutation in 1S enhances sensitivity to activation by the voltage sensor and by caffeine (65).
To test for a role of DHPRs in situ we used mdg myotubes, which form T-SR junctions (20) lacking DHPRs in the T tubule membrane but are structurally normal otherwise. The location of Ca2+ sparks was random in these cells, indicating that T tubules without DHPRs no longer prevented spontaneous opening of the release channels. The DHPR is therefore required for this inhibitory effect.
The test required changes in conditions that made the comparison less clear cut. In dysgenic cells it was necessary to increase [Ca2+]o radically and apply 1 mM caffeine to even observe sparks. Complicating matters, when wild-type cells were observed in the high-[Ca2+] solution with caffeine, an even higher frequency of sparks was induced, together with the loss of the characteristic pattern of locations.
In an attempt to compare wild-type and mdg cells under similar conditions of load, the normal myotubes were imaged in high Ca2+, the addition of 50 µM nifedipine to block Ca2+ entry through L-type Ca2+ channels. The similarity in morphological parameters of Ca2+ sparks recorded in mdg and in normal cells under these conditions was an indication that rough parity in cytosolic and intra-SR Ca2+ levels had been reached. The normal myotubes in higher [Ca2+]o still maintained the suppression pattern, consistent with a necessary role of DHPRs in the basal inhibition of release channels.
We initially thought that the reason for the absence of sparks in mdg cells could be a lower [Ca2+]cyto caused by decreased Ca2+ influx due to the absence of DHPRs (the L-type Ca2+ channels in the plasmalemma), a lower load of Ca2+ in the SR secondary to the diminished influx, or both. Indeed, when [Ca2+]o was increased, the dysgenic cells generated sparks at a higher rate, comparable with the wild-type cells in Krebs.
However, direct measurements of [Ca2+]cyto and [Ca2+]SR did not support this explanation because [Ca2+]cyto was nearly identical in wild-type and mdg cells immersed in Krebs solution. In any case, the final conditions used for comparing patterns of spark location led to similar [Ca2+]cyto and [Ca2+]SR in both types of cells. The [Ca2+] measurements therefore support an inhibitory role of the DHPR, as they show that the difference in patterns may not be attributed to differences in relevant [Ca2+] under the conditions that made the comparison possible.
The suppression effect in embryonic cells is conserved in adults. As first described by Shirokova et al. (53) the topographic segregation between T tubules and Ca2+ sparks has a two-way functional correlate, which consists of an absence of voltage-operated release in the regions where sparks are observed, plus a pattern of voltage-operated release (where it exists) that is devoid of sparks. The failure of voltage sensors in the mammal to elicit sparks was first shown in adult muscle (52). The picture was recently completed with the finding that the elementary events produced by depolarization in the rat are "embers," resulting from the opening of single RyR channels (11).
Thus it appears that DHPRs and RyRs engage in a peculiar functional relationship. It includes a basal inhibition of sparks, plus a dynamic, voltage-dependent activation of opening accompanied by continued inhibition of sparks even when channels open. This peculiar feature is also present in myotubes, as shown by Shirokova et al. (53). Therefore, the DHPR-RyR interaction at work in the embryonic cells produces fully the "adult" effect.
The fact that the inhibitory effect of the DHPR on sparks is observed in both adult and immature mammalian muscle cells has intriguing implications. We showed above that the inhibition of sparks associated with T-tubular structures (and presumably the DHPRs) occurs even in regions where RyR3 isoforms are present. The inhibitory effect, which could be attributed to the mechanical interaction between DHPRs and RyR1, somehow is capable of inhibiting the other isoform, which is believed not to be under any sort of mechanical interaction with voltage sensors (49). That the effect of T-tubular structures on RyR1 channels also prevents RyR3 channels from causing sparks is difficult to explain, as RyR3 are capable of producing sparks when expressed alone in 1B5 cells (63) or in RyR1-null mouse embryonic muscle cells (9). It suggests an intricate interaction, such that the isoform 3 channels cannot engage in coordinated opening unless the RyR1 channels are also enabled.
DHPRs are not sufficient for basal suppression of Ca2+ sparks. Although the present studies point at an important role of the DHPRs in the suppression of sparks, they also demonstrate that the suppressive effect can be overcome. This occurs when normal myotubes are exposed to a high [Ca2+]o. Chun et al. (8) demonstrated an association between "spontaneous" Ca2+ sparks in myotubes and Ca2+ current through L-type Ca2+ channels. The increased unitary current that presumably crosses these channels in conditions of high [Ca2+]o appears to be sufficient to overcome the basal inhibitory effect.
A second instance, in which the inhibitory effect is relieved to a degree, is the chemical permeabilization of the plasmalemma by brief exposure to the mild detergent saponin. The interaction between voltage sensors and release channels may be altered upon exposure to saponin (32). Alternatively, Shirokova and coworkers (26) proposed that an alteration of mitochondrial function, occurring progressively after saponin treatment, is what determines the production of sparks and overcomes the inhibition by T tubule structures.
Work on mechanically skinned skeletal muscle fibers supports the alternative. After the plasmalemma is peeled, T tubules reseal, restore their transmembrane potential, and maintain voltage-dependent skeletal type EC coupling (30). This implies that in peeled fibers DHPRs must be in a normal functional and structural interaction with Ca2+ release channels. As shown by Kirsch et al. (27) and confirmed with rat muscle in our laboratory (B. S. Launikonis and E. Ríos, unpublished observations), spontaneous sparks occur in these fibers, albeit at a lower rate.
Therefore, the presence of the DHPRs is one of the conditions necessary to prevent the occurrence of Ca2+ sparks in the mammal. It should be noted that the functional state of the DHPR is immaterial for this effect, as the suppression persists in depolarized fibers (hence with voltage-inactivated DHPRs; Refs. 11 and 53) or as shown here, in the presence of a high dose of nifedipine, which drives the DHPRs to the same inactivated state.
The studies of Murayama and Ogawa (37, 38) identified an inhibition, described as stabilization against CICR activity, affecting specifically RyRs in the frog and RyR1 in the mammal. This stabilization was not correlated in any detectable degree with a T-SR interaction. The same is true of the inhibition reported by Zhou et al. (67) in muscle of both mammals and amphibians, which they ascribed tentatively to interactions with SR-luminal structures, perhaps calsequestrin. While these effects, and interactions with FKBP12, calsequestrin or other molecules, may be important determinants of the function in situ, they cannot explain the topographic segregation of T tubules and spark generation. Several layers of control are likely to be operating in the native system, reminiscent of those described elsewhere within supramolecular arrays of proteins (12), which endow the participating molecules with properties that can substantially diverge from, and are more subtly controlled than, those observed for the same molecules in isolation.
In conclusion, there are now many evidences for an inhibitory effect of T tubule-associated structures on Ca2+ release channels, which leads to an effective suppression of Ca2+ sparks in situ. The evidence indicates that the RyR1 isoform (or ) is the primary recipient of the inhibition, but the RyR3 isoform is also affected. While DHPRs are needed, they are not sufficient for the inhibitory effect. Many perturbations of the supramolecular arrangement (saponin, mechanical peeling, and mitochondrial alteration) or an increase in the inward current through L-type channels are capable of overcoming the functional inhibition.
Measuring [Ca2+]cyto with SEER. By increasing the dynamic range of ratiometric measurements, SEER makes it possible to image [Ca2+] inside organelles, as initially demonstrated using mag-indo-1 (31). Here we used the higher-affinity dye indo-1 to monitor [Ca2+] in the cytosol. Like other measurements with dyes loaded into the cytosol in AM form, the present one is compromised by entry of dye into organelles. In the APPENDIX, we introduce an approach to correct such errors, which results in Eq. 2 relating [Ca2+] to the measured ratio. Equation 2 is isomorphic with the conventional Eq. 1, but has an effective dissociation constant, Keff, smaller than KD, and an effective minimum ratio, R0, greater than Rmin. This correction is liable to introduce errors, as it assumes the presence of a constant fraction of the dye in the organelles. However, the appendix shows that the magnitude of the correction is proportional to the combined fractional volume of SR and mitochondria in myotubes, which is probably <10% of the total (C. Franzini-Armstrong, unpublished observations). Therefore, even if the dye load and volume of organelles change from cell to cell, the errors in Eq. 2 will probably stay small.
The normal myotubes studied here had on average a [Ca2+]cyto of 152 nM. This value is in good agreement with results obtained in normal myotubes by Shimahara et al. (50) using fura-2, and those measured by Pérez et al. (45) in 1B5 myotubes expressing RyR1 in a careful study with Ca2+-sensitive microelectrodes, but greater than values of Avila et al. (1) using a null-point calibration approach in normal myotubes with indo-1. Considering the many sources of error, the agreement among all these approaches appears reasonable.
[Ca2+]cyto measured in mdg myotubes was on average 172 nM, which is very close to the value in wild-type cells. Similar concentrations in these two types of cells were also reported by Shimahara et al. (50). While we know of no other comparisons between normal and dysgenic myotubes, Weiss et al. (65) reported that uninjected and DHPR 1s-expressing mdg cells had the same resting indo-1 fluorescence ratios. In all, different approaches lead to the conclusion that lack of the L-type Ca2+ channel/voltage sensor per se does not determine a major change in resting cytosolic [Ca2+].
Cytosolic [Ca2+] increased significantly in normal myotubes exposed to high [Ca2+]o. The increase was due in large part to flux through a DHP-sensitive pathway, as it was nearly abolished by 50 µM nifedipine. In dysgenic cultures however, the exposure to high [Ca2+]o did not lead to higher [Ca2+]cyto. One may conclude from these observations that exposure to a high [Ca2+]o increases [Ca2+]cyto through an increase in basal entry of Ca2+ through L-type Ca2+ channels open at rest.
Imaging [Ca2+] in stores of normal and dysgenic myotubes. The present study includes for the first time confocal ratiometric images of a [Ca2+]-sensitive dye in stores of myotubes. Unlike the images of indo-1 in the cytosol, those of mag-indo-1 in organelles reveal structural detail. As the ratiometric measurement is independent, to a good degree, of local dye concentration and volume of the stained organelle, the local variations in the ratio reflect changes in organellar [Ca2+].
In adult cells, two types of structures were described, with clearly different SEER ratios: areas of high ratio, corresponding to SR terminal cisternae, and longitudinal structures of low ratio, identified as mitochondria (31). As shown, for example, in Figs. 9 and 10, there is no comparable pattern in myotubes. It is clear that nuclei do not load well with dye. But perinuclear areas sometimes exhibit a different ratio (Figs. 9C and 10C). Perhaps relative development of mitochondria and SR change among different cultures. Given this variability, we have not attempted functional and structural separation of the main organelles in these cells.
The resting [Ca2+] in stores was 82 µM in normal myotubes. This is substantially lower than in adult frog muscle, where the same technique yields an average of 470 µM for permeabilized fibers equilibrated in a solution with 100 nM [Ca2+]cyto (31). The measurement in myotubes may have been distorted by a contribution from mitochondria. In addition, the concentration of intra-organellar dye was two- to fourfold lower in myotubes than in adult cells, which may have biased the ratio toward lower values. In conclusion, the present measurements, which indicate a major difference in SR load between embryonic mouse and adult frog cells, may have underestimated [Ca2+]SR to some extent.
The measurement, however, should provide a good comparison tool. Thus, it demonstrated an increase in organellar [Ca2+], partially prevented by nifedipine, upon exposing normal myotubes to elevated [Ca2+]o. This result is consistent with the observation of higher [Ca2+]cyto under the same conditions (Fig. 8). The fact that the increase in [Ca2+]cyto took many minutes, even in [Ca2+]o as high as 26 mM, is consistent with the observation that Ca2+ load in stores of embryonic muscle fibers did not change measurably after 10 min of exposure to 8 mM [Ca2+]o (8).
[Ca2+]SR was expected to be close to normal in mdg cells, as many structure-function studies have demonstrated (and relied upon) the integrity of the Ca2+ store in these cells. Surprisingly, intra-organellar [Ca2+] was significantly greater in the dysgenic cells, by about twofold. The present study is not the first to imply greater load in dysgenic cells; Weiss et al. (65) found that the maximal cytosolic Ca2+ transient in response to caffeine was greater in uninjected than in -expressing dysgenic cells, by nearly 40%. As we have shown, dysgenic cells do not produce spontaneous Ca2+ sparks in Krebs, but normal myotubes do. The difference in [Ca2+]SR, could be in part due to the greater leak constituted by large and frequent release events in normal cells at rest.
This explanation may help justify the paradoxical effect of exposure to high [Ca2+]o in dysgenic cells: a decrease in [Ca2+]SR. As shown previously, elevated [Ca2+]o does not significantly increase [Ca2+]cyto in these cells, perhaps because their only pathways of Ca2+ entry may be SOC or ECCE channels (7, 44), with currents that saturate at relatively low [Ca2+]o (7). High [Ca2+]o causes sparks to appear, providing a leak that may shift the steady [Ca2+]SR to a lower value. However, these explanations are incomplete, as they do not provide a reason for the absence of Ca2+ sparks in mdg myotubes at rest, or their appearance in high [Ca2+]o.
The conditions found appropriate for comparing the patterns of spark generation in normal and dysgenic myotubes included high [Ca2+]o, 1 mM caffeine, and, in normal myotubes, 50 µM nifedipine. Under those conditions, free [Ca2+]cyto was 192 nM in the wild-type cells and 118 nM in the dysgenic cells (Fig. 8). In similar solutions, organellar [Ca2+] was 144 µM in both types of cells (Fig. 11). Given these results, the characteristic spatial location of Ca2+ sparks in the wild type is not likely to be due to differences in modulation by Ca2+. It may reflect instead a direct inhibitory effect of the DHPR, missing in dysgenic cells.
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APPENDIX |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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Organelles occupy a certain fraction of the cell volume, and load dye, free or bound, at concentrations that may or may not be equal to that in the cytosol. Let v represent the fraction of the dye load present in organelles (equal to the fractional volume where the organellar dye would partition, if its concentration was the same in organelles and cytosol). Assuming that luminal [Ca2+] is sufficient to saturate the dye, fluorescence F11 will satisfy
 | (3) |
 | (4) |
is f22/f22,Ca, Rmax is f11.Ca/f22,Ca, and Rmin is f11/f22.
This is the equation of Grynkiewycz et al. (25) (Eq. 1 in METHODS), modified to correct for the interference of a dye in organelles. It can be shown that Eqs. 4 and 1 are isomorphic, by defining
 | (5) |
 | (6) |
 | (7) |
(1 – v) and v]. The modified parameters, determined in an on-cell calibration, gave Keff = 394 nM and R0 = 0.51.
The limitations of this correction should be obvious: it assumes the presence of a constant fraction of the dye in the organelles, and it assumes, perhaps more safely given the high affinity of indo-1, that [Ca2+] there is sufficient to saturate the dye. While the first assumption is unlikely to hold as more than a rough approximation, the correction is small. The combined volume of SR and mitochondria in myotubes is probably <10% of the total (C. Franzini-Armstrong,unpublished observations). If v is 0.1, then Keff will be 0.9 KD and R0 will be very close to Rmin (30 times closer to Rmin than Rmax, according to Eq. 6). Therefore, even if the dye load and volume of organelles change from cell to cell, the errors in Eq. 7 (see Eq. 2) will probably remain low.
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GRANTS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONAPPENDIXGRANTSREFERENCES |
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ACKNOWLEDGMENTS |
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Present address for A. González: Fundación Instituto de Estudios Avanzados, Caracas, Venezuela.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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