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. 2003 Jan 24;547(Pt 2):453–462. doi: 10.1113/jphysiol.2002.036129

Metabolic Regulation of Ca2+ Release in Permeabilized Mammalian Skeletal Muscle Fibres

Elena V Isaevaand 1, Natalia Shirokova 1
PMCID: PMC2342647  PMID: 12562922

Abstract

In the present study, the link between cellular metabolism and Ca2+ signalling was investigated in permeabilized mammalian skeletal muscle. Spontaneous events of Ca2+ release from the sarcoplasmic reticulum were detected with fluo-3 and confocal scanning microscopy. Mitochondrial functions were monitored by measuring local changes in mitochondrial membrane potential (with the potential-sensitive dye tetramethylrhodamine ethyl ester) and in mitochondrial [Ca2+] (with the Ca2+ indicator mag-rhod-2). Digital fluorescence imaging microscopy was used to quantify changes in the mitochondrial autofluorescence of NAD(P)H. When fibres were immersed in a solution without mitochondrial substrates, Ca2+ release events were readily observed. The addition of l-glutamate or pyruvate reversibly decreased the frequency of Ca2+ release events and increased mitochondrial membrane potential and NAD(P)H production. Application of various mitochondrial inhibitors led to the loss of mitochondrial [Ca2+] and promoted spontaneous Ca2+ release from the sarcoplasmic reticulum. In many cases, the increase in the frequency of Ca2+ release events was not accompanied by a rise in global [Ca2+]i. Our results suggest that mitochondria exert a negative control over Ca2+ signalling in skeletal muscle by buffering Ca2+ near Ca2+ release channels.

In skeletal muscle, an action potential triggers the release of Ca2+ ions from the sarcoplasmic reticulum (SR) and initiates subsequent contraction. The dihydropyridine receptors (DHPrs) in the transverse tubular system sense membrane depolarization and then through mechanical coupling activate adjacent Ca2+ release channels (ryanodine receptors, Ryrs) in the apposed SR membrane (Schneider & Chandler, 1973; Ríos et al. 1993; Nakai et al. 1996). The initial Ca2+ transient is further amplified by Ca2+-induced Ca2+ release (CICR; Endo et al. 1970; Ford & Podolsky, 1970; Fabiato, 1984). However, it is unclear to what degree CICR contributes to excitation-contraction coupling under physiological conditions (reviewed by Lamb, 2000).

At the subcellular level, CICR is resolved as Ca2+ sparks, which were first detected in confocal microscope images of cardiac myocytes as brief, spatially confined elevations of cytosolic [Ca2+] (Cheng et al. 1993). These events appear to represent the localized release of Ca2+ from a small cluster of Ryrs. Ca2+ sparks were also found in a variety of tissues, including smooth muscle (Nelson et al. 1995), amphibian skeletal muscle (Tsugorka et al. 1995; Klein et al. 1996), embryonic mammalian skeletal muscle and skeletal muscle myotubes (Gÿorke & Gÿorke, 1996; Shirokova et al. 1998; Conklin et al. 1999). A second form of local Ca2+ release was discovered in amphibians (Shirokova & Ríos, 1997). It was termed ‘small event’ Ca2+ release, since the events were smaller than Ca2+ sparks. This form was prominent under experimental conditions that reduced CICR. We proposed that direct interaction between DHPrs and Ryrs gives rise to the small event Ca2+ release, which, in turn, triggers Ca2+ sparks. The idea was supported by the discovery of embers, low-intensity prolongations of Ca2+ sparks elicited by depolarization, in frog skeletal muscle (Gonzalez et al. 2000).

Ca2+ sparks are rarely observed in intact adult mammalian skeletal muscle cells (Conklin et al. 1999). In cut mammalian skeletal muscle fibres, depolarization produced a small event Ca2+ release with no hint of Ca2+ sparks, leading to the suggestion that DHPrs tightly control Ryrs in mammals and prevent CICR (Shirokova et al. 1998). However, Ca2+ sparks were detected recently in skinned adult mammalian muscle fibres (Kirsch et al. 2001), suggesting that CICR does occur under some experimental conditions. The report of Kirsch et al. (2001) stimulated the search for mechanisms that inhibit CICR in intact cells and that may be altered during the permeabilization procedure.

Intracellular metabolic pathways coupled to mitochondria are likely to be disrupted after perforation of the sarcolemmal membrane and subsequent washout of the cytosol. Evidence from a variety of cell types indicates that mitochondria play an important role in Ca2+ homeostasis (for reviews see Babcock & Hille, 1998; Duchen, 1999; Rizutto et al. 2000). In particular, mitochondria serve as a Ca2+ sink at times of Ca2+ excess in the cytoplasm, thus modulating intracellular Ca2+ signals (for reviews see Gunter et al. 1998, 2000). Mitochondria were also shown to affect the spatiotemporal pattern of local Ca2+ signals in smooth (Gordienko et al. 2001) and cardiac (Pacher et al. 2002) muscle myocytes, in Xenopus oocytes (Marchant et al. 2002) and in other tissues. A tight apposition of the organelles with SR membranes facilitates a functional exchange between Ca2+ release from the internal depot and mitochondrial Ca2+ uptake (for reviews see Hajnóczky et al. 2000; Csordás et al. 2001).

Skeletal muscle fibres are rich in mitochondria. Morphological studies have revealed the close proximity of the SR to mitochondria (Ogata &Yamasaki, 1985). This suggests that mitochondria can participate in the regulation of intracellular Ca2+ signals in skeletal muscle. However, to date, our knowledge about functional crosstalk between the two organelles in this tissue is very limited.

The present study was designed to evaluate the link between muscle metabolism and local Ca2+ signalling in skeletal muscle. Our results provide evidence that mitochondria play a substantial role in the regulation of spontaneous Ca2+ release in permeabilized skeletal muscle cells.

METHODS

Cell preparation and solutions

Experiments were carried out on cut skeletal muscle fibres from the extensor digitorum longus (EDL) muscle of the rat. According to the protocol approved by the Institutional Animal Care and Use Committee, female rats (Rattus norvegicus, Sprague-Dawley, 175–200 g) were killed by cervical dislocation under deep anaesthesia induced by intraperitoneal injection of sodium pentobarbital (100–200 mg (kg body weight)−1). Fibres were dissected as described by García & Schneider (1993) and transferred to an experimental chamber similar to that of Ríos et al. (1999). Cells were permeabilized by a 40 s exposure to 50 μg ml−1 saponin and then immersed into one of the ‘internal’ solutions. According to Launikonis & Stephenson (1997), this permeabilization procedure should not substantially affect the Ca2+ loading ability of the SR.

The l-glutamate-based ‘internal’ solution contained (mm): 140 potassium l-glutamate, 10 Hepes, 0.5 EGTA, 5 phosphocreatine (di-Tris salt), 3 Mg-ATP (free [Mg2+]∼ 380 μm) and 0.114 CaCl2 (free [Ca2+]∼ 100 nm). The d-glutamate-based solution contained 140 mm potassium d-glutamate, instead of potassium l-glutamate. In some experiments, 20 or 60 mm d-glutamate was replaced with the corresponding amount of l-glutamate or pyruvate. The free [Ca2+] and [Mg2+] at given total Ca2+, Mg2+, ATP, EGTA and glutamate concentrations were calculated using WINMAX software (Stanford University, CA, USA). Dissociation constants were taken from NIST Critically Selected Stability Constants of Metal Complexes database (A.E. Martell). We assumed that the constants are the same for d and l forms of glutamate. All solutions were titrated to pH 7.0 with KOH and had an osmolarity of ∼300 mosmol kg−1.

Local Ca2+ transients were detected with fluo-3 (50 μm) in the internal solution. Mitochondrial membrane potential (Δψm) was recorded with 10 nm tetramethylrhodamine ethyl ester (TMRE). To monitor mitochondrial matrix [Ca2+] ([Ca2+]m), intact cells were first loaded with 1 μm mag-rhod-2 AM for 10 min at room temperature, then washed and permeabilized as described above. Some cells were co-loaded with MitoTracker Green FM (200 nm). This dye binds to the inner mitochondrial membrane and fluoresces independently of [Ca2+]m or Δψm, thus providing a test of mitochondrial localization of mag-rhod-2 and TMRE. All measurements were performed at room temperature, 30 min after cells were permeabilized.

Fluo-3 was obtained from Biotium (Hayard, CA, USA); TMRE, mag-rhod-2 and MitoTracker Green FM were from Molecular Probes (Eugene, OR, USA); Ru360 was from Calbiochem (La Jolla, CA, USA). All other chemicals were from Sigma (St. Louis, MO, USA).

Fluorescence imaging

Cytoplasmic Ca2+ transients, as well as changes in [Ca2+]m and Δψm, were monitored with a Radiance 2000 confocal scanner (Bio-Rad, Hercules, CA, USA) coupled to a Zeiss Axiovert 100 microscope with a × 40, 1.2 NA, water-immersion objective (Zeiss, Oberkohen, Germany). Fluo-3 and MitoTracker Green FM were excited at 488 nm with an Argon laser. TMRE and mag-rhod-2 were excited with a HeNe laser at 543 nm. The emitted light was collected above 500 nm (fluo-3) and above 570 nm (TMRE and mag-rhod-2). In co-labelling experiments, cells were simultaneously excited at 488 and 543 nm with the Argon and HeNe lasers. The beam of emitted light was split at 560 nm, and the two resulting beams were collected simultaneously with two detectors equipped with a 515 ± 30 nm band-pass filter and a 600 nm long-pass emission filter, respectively. Fibres were imaged in full-frame (XY) mode at 500 lines s−1. Unless otherwise specified, series of 40 (102.8 × 102.8 μm) images were acquired at 0.2–0.33 Hz at randomly selected locations on each fibre. Some linescan (X-t) images were also obtained with 512 pixels (0.2 μm pixel−1) in the x direction and 1000 pixels (2 ms line−1) in the t direction. These images revealed a complex morphology of spontaneous events of Ca2+ release (data not shown) similar to that reported previously by Kirsch et al. (2001).

Discrete events of Ca2+ release were identified with an automatic detection method (Cheng et al. 1999) modified for localization of events in XY images. Images of fluorescence F(x,t) were first normalized to the resting fluorescence F0(x,t) using a procedure similar to that described by Brum et al. (2000). Then, the automatic detector identified event areas with normalized fluorescence F/F0 greater than the threshold. The threshold was usually set to be m+ 3σ, where m and σ are the mean and standard deviation of the resting fluorescence, respectively. Because XY images provide little information about the event morphology, we will term all events as Ca2+ sparks.

NAD(P)H fluorescence was imaged with CH1 CCD (Photometrics, USA), as described previously by Hajnóczky et al. (1999). Fibres were excited at 360 nm and the light emitted above 420 nm was recorded. A single series of 300 images at 0.33 Hz was acquired in each experiment.

Statistics

All values are presented as means ±s.e.m. Student's t test for paired observations was used to assess significance. Values were considered significant if P < 0.05.

RESULTS

Mitochondrial substrates inhibit spontaneous events of Ca2+ release

In the first group of experiments, we compared the frequencies of spontaneous events of Ca2+ release in fibres incubated in solutions containing different amounts of mitochondrial substrates. While l-glutamate and pyruvate are substrates of the tricarboxylic acid cycle, d-glutamate is not. Figure 1 represents data from an experiment in which a fibre was sequentially incubated in d-glutamate- and l-glutamate-based internal solutions. Figure 1A shows fluorescence images obtained 30 min after the fibre was immersed into the control, d-glutamate-based, internal solution (a), 10 min after changing the internal solution to that based on l-glutamate (b) and 10 min after returning to the control solution (c). Sets of 40 sequential images were acquired in each experimental condition. The sparks were identified in each image of the set and are presented in Fig. 1B as accumulative masks. The spark frequency (f) was determined as the mean number of events per unit area and time. When the fibre was immersed in the d-glutamate-based solution, Ca2+ sparks were readily detected (f= 6.2× 10−4μm−2 s−1). When d-glutamate in the internal solution was replaced with l-glutamate, Ca2+ sparks were substantially suppressed (f= 0.8× 10−4μm−2 s−1). The inhibitory effect of l-glutamate was reversed after fibre washout with d-glutamate (f= 5.2× 10−4μm−2 s−1).

Figure 1. Ca2+ sparks in l- and d-glutamate.

Figure 1

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A, fluorescence images of a cell immersed in internal solutions based on d-glutamate (a) and l-glutamate (b), and after washout (c). B, corresponding accumulative binary images for the regions with F/F0 > 3 s.d. Cell no. 021102–3.

Figure 2A summarises the effect of l-glutamate on the frequency of Ca2+ sparks in 18 fibres. The frequency of sparks varied considerably in the control condition (f= (3.14 ± 0.43) × 10−4μm−2 s−1). However, l-glutamate reduced the number of detected events in all fibres studied (f= (0.55 ± 0.12) × 10−4μm−2 s−1). The inhibitory effect was statistically significant, as determined by paired Student's t test (P= 2.6× 10−5). The effect was also partially reversible (f= (1.83 ± 0.29) × 10−4μm−2 s−1).

Figure 2. Effects of oxidizable mitochondrial substrates on the frequency of Ca2+ sparks.

Figure 2

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A, comparison of the frequency of Ca2+ sparks in d-glutamate (d-glut) and l-glutamate (l-glut) internal solutions (n= 18). B, frequency of Ca2+ sparks in d-glutamate solution, and after d-glutamate was replaced with 20 mm (n= 12) or 60 mm (n= 16) l-glutamate, or with 20 mm pyruvate (n= 6). Box plots are used to show the details of the distribution of frequencies of Ca2+ sparks under each experimental condition. The boundaries of each box indicate the 25th and 75th percentiles, whiskers indicate the 90th and 10th percentiles, the line within the box marks the median frequency and the asterisk represents the mean frequency.

Changes in the frequency of Ca2+ sparks did not correlate with the variations in the resting fluorescence. In the experiments summarized in Fig. 2A, the resting fluorescence increased monotonically by 6 ± 2 % (n= 18) during the experiment (∼60 min). This increase was comparable with that observed in the control group of experiments, where no changes of the internal solution were performed (7 ± 2 %, n= 11).

The frequency of Ca2+ sparks also decreased upon partial replacement of d-glutamate in the internal solution with l-glutamate or pyruvate. Figure 2B summarises the results of the experiments where 20 or 60 mm d-glutamate was substituted with l-glutamate, or 20 mm d-glutamate was replaced with pyruvate. On average, 20 mm l-glutamate reduced the frequency of sparks by 39 ± 7 % (n= 12, P= 4.5× 10−3), 60 mm l-glutamate decreased it by 61 ± 7 % (n= 16, P= 5.1× 10−5) and 20 mm pyruvate inhibited Ca2+ sparks by 94 ± 6 % (n= 6, P= 1.4× 10−3).

Mitochondrial substrates stimulate mitochondrial functions

In order to investigate the effects of mitochondrial substrates on mitochondrial function, we monitored the mitochondrial membrane potential and the redox state of the mitochondrial NAD system. Figure 3A illustrates mitochondrial localization of the potentiometric dye TMRE. TMRE fluorescence (Fig. 3Ab) overlapped with MitoTracker Green FM signal (Fig. 3Aa), as indicated by the yellow colour of the overlay image (Fig. 3Ac). Clusters of sarcomere-bound mitochondria, as well as some mitochondria columns, were clearly distinguishable. Figure 3B and C illustrates changes in TMRE fluorescence when d-glutamate was replaced with l-glutamate and after washout. Images were acquired every 10 s. TMRE fluorescence was spatially averaged over the region of interest corresponding to the fibre. The averaged values are plotted vs. time on Fig. 3B. TMRE exhibits a Nernstian distribution across the inner mitochondrial membrane, therefore the averaged fluorescence is represented on a logarithmic scale. Since at the concentration used in our experiments (10 nm) the dye is unlikely to be subject to self-quenching, the decrease in TMRE fluorescence indicates a membrane depolarization. The mitochondrial membrane gradually depolarized while the fibre was incubated in d-glutamate. The membrane potential was restored in l-glutamate and decreased again upon washout. As expected, the mitochondrial uncoupler carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP, 2.5 μm), added in combination with oligomycin (2.5 μm), an inhibitor of F1/F0 ATP synthase, rapidly depolarized the mitochondrial membrane. Inserts display images of TMRE fluorescence obtained at selected times during this experiment. The mean values of the averaged TMRE fluorescence were determined from 10 images acquired before the addition of l-glutamate and from 10 images acquired right before its washout with d-glutamate. l-glutamate increased the TMRE fluorescence intensity on average by 63 ± 3.1 % (n= 5, P= 1.4× 10−5).

Figure 3. Effect of l-glutamate on mitochondrial potential.

Figure 3

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A, arrangement of mitochondria in the extensor digitorum longus muscle fibre. a, mitochondria labelled with MitoTracker Green FM. TMRE fluorescence (b) overlapped with the MitoTracker Green FM signal, as indicated by the yellow colour in c. B, changes in mitochondrial potential (Δψm) in response to the addition of l-glutamate, after its washout and after application of mitochondrial uncouplers (FCCP and oligomycin). a.u., arbitrary units. C, confocal images of TMRE fluorescence acquired at the times indicated by the numbers on the plot in B. Cell no. 122101–3.

We also investigated how the substrates alter the redox state of the mitochondrial NAD system by monitoring NAD(P)H autofluorescence. Figure 4 illustrates changes in the spatially averaged NAD(P)H fluorescence signal. The mean values of the averaged NAD(P)H fluorescence were determined, as described above, in d- and l-glutamate. In this, and seven other experiments, l-glutamate reversibly increased the NAD(P)H signal by 43 ± 8.2 % (P= 1.7× 10−3). Thus, in our preparation, like in mouse quadriceps muscle (Kuznetsov et al. 1998), mitochondrial substrates enhanced the NAD(P)H production.

Figure 4. Effect of l-glutamate on the mitochondrial NAD system.

Figure 4

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This plot shows changes in NAD(P)H autofluorescence after addition of the substrate and following its washout. Cell no. 112101–11.

Mitochondrial inhibitors promote Ca2+ sparks in permeabilized fibres

Mitochondrial Ca2+ uptake operates via an electrogenic uniporter (for review see Gunter et al. 1998). Interventions that reduce the mitochondrial membrane potential (e.g. FCCP, antimycin A), or block the mitochondrial uniporter (e.g. Ru360), inhibit mitochondrial Ca2+ uptake. We studied how these drugs affect the frequency of Ca2+ release events. All drugs were added to the l-glutamate-based solution.

Figure 5A represents fluorescence images of a fibre in the l-glutamate (control) solution (a) and after 20 μm Ru360 was added (b). Panels c and d show accumulative masks obtained from sets of 40 images acquired under each condition. In this experiment, the frequency of Ca2+ sparks increased from 0.50 × 10−4 to 3.16 × 10−4μm−2 s−1 after 10 min of incubation with Ru360. Figure 6A summarizes the results of nine similar experiments. In each fibre, the frequency of sparks was determined under control conditions, as well as 5 and 10 min after Ru360 was applied. On average, the spark frequency increased by 229 ± 54 % (P= 2.5× 10−3) after 10 min of fibre incubation with Ru360. The corresponding increase in the level of resting fluorescence was very small (1.80 ± 0.01 %). Application of Ru360 to the fibres incubated in d-glutamate solution did not produce a significant increase in the Ca2+ spark frequency (n= 5, P= 0.16, data are not shown).

Figure 5. Mitochondrial inhibitors promote Ca2+ sparks.

Figure 5

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A, fluorescence images of a cell in l-glutamate (control) solution (a) and after the addition of Ru360 (b). c and d, accumulative masks of spark regions. Cell no. 042502–6. B and C, effects of antimycin A (cell no. 061302–2) and FCCP with oligomycin (cell no. 050902–1).

Figure 6. Effects of mitochondrial inhibitors on the frequency of Ca2+ sparks.

Figure 6

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A, frequencies of Ca2+ sparks under control conditions (black bar) and after Ru360 was added (open bars), n= 9. B, C and D, effects of antimycin A (n= 8), FCCP with oligomycin (n= 8) and oligomycin (n= 11), respectively.

The respiratory chain (complex III) inhibitor antimycin A (2.5 μm) also promoted Ca2+ sparks (Fig. 5B and Fig. 6B). After 5 min of fibre incubation with antimycin A, spark frequency increased on average by 184 ± 12 % (n= 8, P= 0.017). In this case, however, the increase in Ca2+ spark frequency was accompanied by noticeable increase in the resting fluorescence (11.5 ± 2.5 %). This increase was substantially larger than that observed under control conditions (see above) and is likely to reflect the increase in cytoplasmic [Ca2+] following the application of antimycin A.

FCCP (2.5 μm), applied in combination with oligomycin (2.5 μm), increased the frequency of Ca2+ sparks on average by 331 ± 115 % (n= 8, P= 0.013). The effect on Ca2+ sparks was rapid and transient (Fig. 5C and Fig. 6C). Ca2+ sparks disappeared after 10 min incubation with FCCP and oligomycin, probably due to depletion of Ca2+ from the SR (Arnaudeau et al. 2001). The initial increase in the frequency of Ca2+ sparks was accompanied by substantial changes in resting fluorescence, which increased by 22.2 ± 3.9 % within 2 min after the drugs were applied. Oligomycin (2.5 μm, n= 11), applied separately, failed to promote Ca2+ sparks (Fig. 6D).

Thus, pharmacological interventions that disrupted mitochondrial Ca2+ uptake promoted spontaneous Ca2+ release in permeabilized skeletal muscle fibres.

Mitochondrial [Ca2+] signals

The experiments represented in Fig. 7 were designed to measure changes in [Ca2+]m after the application of mitochondrial inhibitors. Figure 7A demonstrates mitochondrial localization of mag-rhod-2. Mag-rhod-2 fluorescence spatially overlapped with the signal of MitoTracker Green FM, as indicated by the yellow colour. The staining pattern was similar to that obtained with TMRE (Fig. 3A). A substantial amount of Ca2+ was trapped inside the mitochondria under the conditions of our experiments; because of this, we used low-affinity mitochondrial Ca2+ indicator mag-rhod-2 for measurements of [Ca2+]m. It is possible that the absence of Na+ in our internal solutions inhibited the efflux of Ca2+ from the organelles. However, a significant mitochondrial Ca2+ content was also measured by Lännergren et al. (2001) in intact frog and mouse skeletal muscle fibres under resting conditions.

Figure 7. Effects of mitochondrial inhibitors on [Ca2+]m.

Figure 7

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A, overlay of MitoTracker Green FM and mag-rhod-2 fluorescence. B, changes in [Ca2+]m following the addition of Ru360 (cell no. 053020–5), antimycin A (cell no. 071502–1) and FCCP with oligomycin (cell no. 052902–4). The arrow indicates the time when drugs were applied. C, summary of the effects of the inhibitors. D, images of mag-rhod-2 fluorescence in control conditions and after the application of Ru360 (top) and FCCP with oligomycin (bottom).

Figure 7B represents spatially averaged mag-rhod-2 fluorescence, which was normalized to the average fluorescence calculated from 10 images acquired before the drugs were applied. In many experiments, application of mitochondrial inhibitors caused loss of the mitochondrial Ca2+. However, each inhibitor acted somewhat differently.

In some fibres, application of Ru360 caused a slight decrease in [Ca2+]m (Fig. 7B and D). Overall, this decrease was not statistically significant (P= 0.076, n= 11). In addition, we did not detect any significant decrease in [Ca2+]m after removal of the mitochondrial substrates from the internal solution (n= 15, P= 0.096). The loss of the mitochondrial Ca2+ was rapid when FCCP was applied together with oligomycin. The mag-rhod-2 fluorescence decreased by 87 ± 4 % (n= 8, P= 2.1× 10−7) within 3 min after application of the drugs. Corresponding images of mag-rhod-2-loaded fibres are presented in Fig. 7D. Application of antimycin A caused an intermediate effect: mag-rhod-2 fluorescence declined by 36 ± 9 % (n= 5, P= 0.018) after 10 min of incubation with antimycin A.

DISCUSSION

As mentioned in the introduction, discrete Ca2+ release events have been found recently in skinned mammalian skeletal muscle fibres (Kirsch et al. 2001). By analogy with amphibian skeletal muscle, the events were roughly divided into two groups: large, brief sparks, or small, long-lasting embers. The mechanisms that gate these events and modulate their frequency are poorly understood. Our knowledge about factors that prevent the appearance of Ca2+ sparks in intact mammalian muscle is also very limited. The aim of this article was to evaluate whether spontaneous Ca2+ signals in mammalian skeletal muscle fibres are regulated by the metabolic state of the fibres.

Our main finding is that the frequency of Ca2+ release events in permeabilized mammalian skeletal muscle depends greatly on mitochondrial respiration. Stimulation of mitochondrial activity by the addition of mitochondrial substrates resulted in a reversible inhibition of Ca2+ sparks. Moreover, Ca2+ spark activity was enhanced by drugs that are known to suppress mitochondrial respiration (antimycin A), to collapse mitochondrial potential (FCCP and antimycin A), or to directly inhibit mitochondrial Ca2+ uptake (Ru360).

Localization of mitochondria in skeletal muscle fibres

Mitochondria support oxidative metabolism, and the volume of mitochondria and surface area of mitochondrial cristae can be an index of a cell's oxidative capacity. Muscles that are composed mostly of slow-oxidative fibres (e.g. the rat soleus muscle) have higher volumes of mitochondria per unit muscle volume (∼10 %), and muscles that contain mostly fast-glycolytic fibres (e.g. rat gastrocnemius muscle) have lower mitochondria volume densities (∼2 %; reviewed by Eisenberg, 1983). The EDL skeletal muscle of rat contains about 60 % fast-oxidative-glycolytic (FOG) fibres (Ariano et al. 1973). FOG fibres have high glycolytic capacity along with high mitochondrial content (∼7 %, Eisenberg, 1983).

In mammalian skeletal muscle fibres, most mitochondria are found packed in between the myofibrils. Large numbers of mitochondria are also found in parallel clusters under the sarcolemma. The distribution and amount of mitochondria and the SR are constant through the whole length of the fibre. Scanning electron microscopy has revealed that I-band-limited mitochondria are adjacent to the terminal cisternae of the SR (Ogata & Yamasaki, 1985). The close proximity of these two organelles provides the structural basis for their functional interaction. In FOG fibres, some mitochondria are also arranged into longitudinal columns (e.g. Fig. 3A). The columns extend over several sarcomeres and often approach the SR network, providing an additional possibility for crosstalk between the two organelles.

Mitochondrial Ca2+ uptake in skeletal muscle

Studies with mitochondria isolated from various skeletal muscles have shown that these organelles are capable of sequestering Ca2+ (Sembrowich et al. 1985; Madsen et al. 1996). Direct measurements of mitochondrial [Ca2+] in skeletal muscle myotubes with the Ca2+-sensitive photoprotein aquorin targeted to mitochondria, revealed large mitochondrial Ca2+ transients in response to depolarization or application of Ryr agonists (Challet et al. 2001; Robert et al. 2001). It has also been shown that mitochondria promote relaxation of mitochondria-rich skinned skeletal muscle fibres (Gillis, 1997). However, the role of mitochondrial Ca2+ uptake in the clearance of cytosolic Ca2+ after electrical stimulation in intact skeletal muscle fibres remains unclear. Lännergren et al. (2001) compared mitochondrial Ca2+ signals in intact Xenopus and mouse fast-twitch fibres subjected to tetanic stimulation. It was concluded that while amphibian muscle mitochondria are able to accumulate Ca2+ during repetitive electrical stimulation, mouse mitochondria are not. Further research is needed to clarify the difference.

Mitochondria and local Ca2+ signalling

The data presented here demonstrate that energized mitochondria greatly inhibit the spontaneous release of Ca2+ from the SR in permeabilized mammalian skeletal muscle fibres. Our results are in agreement with recent observations of mitochondria shaping the spatio-temporal pattern of local Ca2+ signals in cardiac (Pacher et al. 2002) and smooth muscle myocytes (Gordienko et al. 2001) and in Xenopus oocytes (Marchant et al. 2002).

Two possible mechanisms by which mitochondria might control Ca2+ sparks should be considered. The first mechanism may be related to the major function of mitochondria - production of ATP. Disruption of ATP production could affect Ryr channel gating and Ca2+ uptake into the SR via the Ca2+ pump. However, this mechanism is unlikely to be responsible for the effects found in our experiments because: (1) cells were superfused with 3 mm ATP to minimize the contribution of metabolically generated ATP; (2) inhibition of ATP production with oligomycin had no long-lasting effects on the frequency of Ca2+ sparks (Fig. 6D).

An alternative mechanism involves the ability of mitochondria to accumulate Ca2+ via the mitochondrial Ca2+ uniporter. Although the Ca2+ uniporter requires relatively high levels of [Ca2+]i in order to be activated (Kd > 1 μm), studies of a variety of tissues have demonstrated that it plays an important role in shaping intracellular Ca2+ signalling under physiological conditions (for reviews see Babcock & Hille, 1998; Duchen, 1999; Rizutto et al. 2000; Csordás et al. 2001). To explain the paradox, it was suggested that mitochondria are strategically positioned in close proximity to Ca2+ release sites (Rizzuto et al. 1998). The latter allows the organelles to sense microdomains of high [Ca2+] during Ca2+ release, thereby promoting the efficiency of Ca2+ uptake via the uniporter. Close apposition of mitochondria and the SR in skeletal muscle is therefore a prerequisite for functional interactions between the organelles.

Under the conditions of our experiments, skeletal muscle mitochondria accumulate a substantial amount of Ca2+. In most cases, [Ca2+]m increased monotonically throughout the duration of the experiment. It is likely that Ca2+ released from the SR is delivered to the mitochondria via the Ca2+ uniporter, as the application of its inhibitors (antimycin A, FCCP in combination with oligomycin, and Ru360) greatly reduced the ability of mitochondria to sequester Ca2+ and, in some cases, led to a decrease in [Ca2+]m. As the removal of Ca2+ from mitochondria is dispatched largely by the mitochondrial Na+-Ca2+ exchanger (reviewed by Bernardi, 1999), the absence of Na+ in our intracellular solutions diminished the ability of mitochondria to extrude Ca2+. After collapse of the mitochondrial membrane potential, Ca2+ is likely to egress the mitochondria via permeability transition pores (Ichas & Mazat, 1998) or the Ca2+ uniporter operating in the reverse mode (Montero et al. 2001). In agreement with the latter possibility, application of the uniporter inhibitor Ru360 did not induce a substantial loss in [Ca2+]m.

Interventions that interfere with mitochondrial Ca2+ uptake stimulated Ca2+ spark activity. The increase in the frequency of Ca2+ sparks could be a consequence of the rise in global [Ca2+]i. Zhou et al. (2002) reported a 4.8-fold increase in event frequency when cytoplasmic [Ca2+] was changed from 100 to 400 nm. An elevation of resting fluorescence (hence an increase in [Ca2+]i) has been reported using fluo-3 in our experiments with antimycin A, and FCCP with oligomycin. In this case, Ca2+ released from the mitochondria after the collapse of Δψm can cause an additional activation of Ryrs via the CICR mechanism.

However, the dramatic increase in spark frequency observed after removal of mitochondrial substrates, or after direct inhibition of the Ca2+ uniporter with Ru360, was not associated with either the increase in resting [Ca2+]i or with the decrease in [Ca2+]m. Hence, the most plausible mechanism for mitochondrial suppression of Ca2+ sparks involves buffering of [Ca2+] in the vicinity of Ca2+ release channels.

Mitochondrial Ca2+ uptake can act in two ways. It can decrease [Ca2+] below the detection threshold during Ca2+ release events and it can prevent the occurrence of events by lowering [Ca2+] near Ryrs. In the first scenario, Ca2+ sparks will permit delivery of Ca2+ directly to the mitochondrial matrix, bypassing fluo-3. In agreement with this possibility, elementary mitochondrial Ca2+ signals (Ca2+ marks) were recently detected in cardiac myotubes (Pacher et al. 2002). In the second scenario, lowering of [Ca2+] in the vicinity of Ryr channels will reduce their open probability and hence inhibit CICR. It is possible that both of these scenarios are involved.

In summary, our results show that Ca2+ spark frequency in skinned skeletal muscle fibres is regulated dynamically by mitochondrial Ca2+ sequestration. We suggest that this process, among others, contributes to the inhibition of CICR in intact mitochondria-rich muscle cells.

Acknowledgments

We thank Drs Laurence Gaspers, John Reeves, Roman Shirokov and Andrew Thomas for helpful discussions and critical reading of the manuscript. This work was supported by grants from the National Institute of Health (R01-AR45690) and Muscular Dystrophy Association.

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