Differential impact of mitochondrial positioning on mitochondrial Ca²⁺ uptake and Ca²⁺ spark suppression in skeletal muscle

Abstract

Muscle contraction requires ATP and Ca²⁺ and, thus, is under direct control of mitochondria and the sarcoplasmic reticulum. During postnatal skeletal muscle maturation, the mitochondrial network exhibits a shift from a longitudinal (“longitudinal mitochondria”) to a mostly transversal orientation as a result of a progressive increase in mitochondrial association with Ca²⁺ release units (CRUs) or triads (“triadic mitochondria”). To determine the physiological implications of this shift in mitochondrial disposition, we used confocal microscopy to monitor activity-dependent changes in myoplasmic (fluo 4) and mitochondrial (rhod 2) Ca²⁺ in single flexor digitorum brevis (FDB) fibers from 1- to 4-mo-old mice. A robust and sustained Ca²⁺ accumulation in triadic mitochondria was triggered by repetitive tetanic stimulation (500 ms, 100 Hz, every 2.5 s) in FDB fibers from 4-mo-old mice. Specifically, mitochondrial rhod 2 fluorescence increased 272 ± 39% after a single tetanus and 412 ± 45% after five tetani and decayed slowly over 10 min following the final tetanus. Similar results were observed in fibers expressing mitochondrial pericam, a mitochondrial-targeted ratiometric Ca²⁺ indicator. Interestingly, sustained mitochondrial Ca²⁺ uptake following repetitive tetanic stimulation was similar for triadic and longitudinal mitochondria in FDB fibers from 1-mo-old mice, and both mitochondrial populations were found by electron microscopy to be continuous and structurally tethered to the sarcoplasmic reticulum. Conversely, the frequency of osmotic shock-induced Ca²⁺ sparks per CRU density decreased threefold (from 3.6 ± 0.2 to 1.2 ± 0.1 events·CRU⁻¹·min⁻¹·100 μm⁻²) during postnatal development in direct linear correspondence (r² = 0.95) to an increase in mitochondrion-CRU pairing. Together, these results indicate that mitochondrion-CRU association promotes Ca²⁺ spark suppression but does not significantly impact mitochondrial Ca²⁺ uptake.

skeletal muscle contraction requires ATP and Ca²⁺ and, thus, is under direct control of two major intracellular organelles, mitochondria and Ca²⁺ release units (CRUs) or triads, formed by the association of two sarcoplasmic reticulum (SR) terminal cisternae flanking a central bisecting transverse tubule (T-tubule). Mitochondria are the powerhouse of the cell, being responsible for aerobic production of cellular ATP required for myosin crossbridge cycling and Ca²⁺ reuptake by sarco(endo)plasmic reticulum Ca²⁺-ATPase (SERCA) pumps (24). CRUs are sites of excitation-contraction coupling, the process that triggers SR Ca²⁺ release and subsequent muscle contraction in response to an action potential propagating down the T-tubule system. Interestingly, in fully mature fast-twitch muscle fibers, the two organelles are in close proximity with each other at the level of the myofibril I band (3, 4, 22, 23, 32).

We recently reported that the association of mitochondria to CRUs occurs progressively during postnatal development in a process that involves an increase in the number of both organelles and their coordinated migration to the I band region during maturation (4). Specifically, in adult fast-twitch muscle fibers [e.g., flexor digitorum brevis (FDB) fibers from 4-mo-old mice], the A band region generally lacks mitochondria (although they are occasionally observed), especially relative to the number of mitochondria observed within the I band. On the other hand, longitudinal mitochondria within the A band region are much more commonly observed in FDB fibers from young mice (e.g., FDB fibers from 0.5- to 1-mo-old mice). This process is mediated by small (∼10 nm) electron-dense linkages, or tethers, that bridge the outer mitochondrial membrane with the SR on the side opposite to the T-tubule. The tethers provide a strong structural link, positioning mitochondria on the side of the SR opposite to Ca²⁺ release channels or type 1 ryanodine receptors (RYR1s) and restrict mitochondrial movement away from sites of SR Ca²⁺ release (4). This intimate, highly coordinated coupling of mitochondria to CRUs during postnatal development is hypothesized to provide a structural basis for localized cross talk between the two organelles (24).

Although this hypothesis has yet to be definitively demonstrated, previous studies have provided evidence for mitochondrial Ca²⁺ uptake (orthograde SR-mitochondrial coupling) during stimulation of adult skeletal muscle fibers in vitro (27), in situ (7, 17, 29), and in vivo (25). Additionally, mitochondrial activity inhibits local Ca²⁺ release events (termed Ca²⁺ spark suppression) from adjacent CRUs (retrograde SR-mitochondrial coupling) (for review see Ref. 24). In their capacity to generate, scavenge, and detoxify reactive oxygen (ROS)/reactive nitrogen species (RNS), CRU-paired mitochondria may create a spatially confined Ca²⁺/ATP/ROS domain that maintains the adjacent CRU and associated proteins (e.g., RYR1) in the proper redox state (13, 14, 19), decreasing susceptibility to Ca²⁺-induced Ca²⁺ release and, thus, Ca²⁺ spark activity. The nature of this bidirectional interaction has been assessed through the use of pharmacological interventions to disrupt mitochondrial function and the local redox control mechanism, to alter the local Ca²⁺ microdomain, and to block mitochondrial Ca²⁺ uptake (7, 14, 18, 19, 29).

Mitochondrial Ca²⁺ uptake enhances aerobic ATP production by stimulating Ca²⁺-dependent dehydrogenases that generate NADH (pyruvate, NAD-isocitrate, and 2-oxoglutarate dehydrogenases) and augmentation of the ATP synthetic capacity of the F₁F₀-ATPase (2, 5, 24). However, since the increase in mitochondrial Ca²⁺ during a single twitch is short-lived (<200 ms) (25) and physiological activation of skeletal muscle usually involves repetitive high-frequency neuromuscular stimulation, we hypothesized that sustained increases in mitochondrial Ca²⁺ occur during repetitive tetanic stimulation and that mitochondrion-CRU pairing is a prerequisite for this phenomenon. To test the impact of intimate mitochondrion-CRU coupling on bidirectional Ca²⁺ signaling between the SR and mitochondria, we took advantage of the fact that mitochondrion positioning changes during postnatal maturation (3, 4). Thus we characterized the degree and properties of mitochondrial Ca²⁺ uptake during tetanic stimulation and Ca²⁺ spark suppression in single skeletal muscle fibers at different times during postnatal maturation, with the expectation that the bidirectional interaction would be strengthened as mitochondrion-CRU pairing progressively increases.

MATERIALS AND METHODS

Muscle fiber preparation.

All animals were housed in a pathogen-free area at the University of Rochester, and experiments were carried out in accordance with procedures reviewed and approved by the local University Committee on Animal Resources. Mice were killed immediately prior to experiments by regulated CO₂ delivery followed by cervical dislocation. Single acutely dissociated FDB muscle fibers were isolated from young (0.5- and 1-mo-old) and adult (2- and 4-mo-old) mice, as described previously (4). All single-fiber experiments were conducted at room temperature (22–23°C).

Myoplasmic Ca²⁺ measurements.

FDB fibers were loaded with 4 μM mag-fluo 4-AM for 25 min at room temperature and then exposed for 20 min to dye-free solution supplemented with 25 μM N-benzyl-p-toluene sulfonamide (BTS) to block contractions (8, 28). A series of repetitive high-frequency tetani (1–40 tetani elicited by 1- to 5-ms, 8-V pulses at 100 Hz for 500 ms delivered at 2.5-s intervals; Fig. 1A) were elicited using an extracellular 200 mM NaCl-filled electrode placed near the fiber of interest. No significant difference was observed in the magnitude of the first evoked Ca²⁺ transient elicited during the tetanus using 1- or 5-ms stimuli [peak relative change in fluorescence (ΔF/F₀) = 0.56 ± 0.03 and 0.50 ± 0.04, n = 14 for 1- and 5-ms stimuli, respectively]. Thus, only single action potentials were elicited, even when 5-ms stimuli were used. For measurements of global myoplasmic Ca²⁺ (Fig. 1A), a rectangular region of interest was selected in mag-fluo 4-loaded fibers and excited at 480 ± 15 nm. Fluorescence emission at 535 ± 20 nm was collected at 10 kHz using a photomultiplier detection system. Data were analyzed offline using Clampfit and SigmaPlot software packages.

Fig. 1.

Myoplasmic and mitochondrial Ca²⁺ transients in flexor digitorum brevis (FDB) fibers during twitch and repetitive tetanic stimulation. A: tetanic stimulation protocol consisting of 1- to 5-ms, 8-V square-wave pulses at 100 Hz for 500 ms (top) delivered at 2.5-s intervals (0.2 duty cycle) for a total of 40 successive tetani (bottom). ▴ indicate rhod 2 image acquisition (after 1, 5, 10, 20, 30, and 40 tetani) shown in Figs. 3 and 5. Inset: increase in myoplasmic mag-fluo 4 fluorescence during the 5th tetanus of the protocol taken from a representative FDB fiber. B: peak mag-fluo 4 fluorescence (ΔF/F₀) during each tetanus in FDB fibers from 1-mo-old mice. Values are means ± SE. C: representative confocal images of MitoTracker Green FM (MTG, top) and rhod 2 fluorescence (middle and bottom) in FDB fibers from a 4-mo-old mouse before and after twitch (1 Hz) or tetanic (100 Hz for 500 ms) stimulation. MTG (top right) and rhod 2 (middle right) fluorescence intensity profiles are shown along the lines of interest marked in the corresponding x-y images. AU, arbitrary units.

Electroporation in FDB muscle.

Mitochondrial-targeted ratiometric pericam (mt-pericam) (21) was electroporated into FDB muscles of adult mice, as described previously (10). Briefly, after anesthesia by intraperitoneal injection of 100 mg/kg ketamine, 10 mg/kg xylazine, and 3 mg/kg acepromazine, bovine hyaluronidase (7 μl/foot, 2 μg/μl) was injected subcutaneously into both hindlimb footpads. After 1 h, 30 μg of mt-pericam cDNA (10 μl total volume in 71 mM NaCl) were similarly injected, and the FDB muscle was electroporated (20-ms, 100 V/cm pulses at 1 Hz for 20 s) using gold-plated electrodes placed perpendicular to the long axis of the muscle, close to the proximal and distal tendons. Individual FDB fibers were isolated 1 wk after electroporation, as described previously (4).

Confocal x-y imaging and analysis of mitochondrial Ca²⁺ accumulation during repetitive tetanic stimulation.

Acutely dissociated FDB fibers were loaded first with 5 μM rhod 2-AM for 30 min at room temperature in Ringer solution (in mM: 146 NaCl, 5 KCl, 1 MgCl, 2 CaCl₂, and 10 HEPES, pH 7.4) supplemented with 25 μM BTS to inhibit cell movement. Minimizing dye concentration and loading time at room temperature facilitated preferential mitochondrial rhod 2 loading. Fibers loaded with rhod 2 were then incubated with 300 nM MitoTracker Green (MTG; Fig. 1C) or 10 μM fluo 4-AM (Fig. 1; see Figs. 2 and 4) and then exposed for 20 min to dye-free solution supplemented with 25 μM BTS. Additional experiments were conducted in fibers loaded with 10 nM tetramethylrhodamine ethyl ester (TMRE), a cationic, mitochondrial-selective fluorescent membrane potential indicator, and 10 μM fluo 4-AM (see Fig. 5). Time series of x-y images (256 × 256 pixels/frame, 0.2 μm/pixel, 690-ms scan duration) were acquired on a confocal microscope (Nikon Eclipse C1 Plus) equipped with a ×40 1.3 numerical aperture (NA) oil objective (SuperFluor) by sequential excitation of green (MTG and fluo 4) and red (rhod 2 and TMRE) fluorophores using 488- and 543-nm lasers, respectively. Repetitive high-frequency tetanic stimulation trains (1–40 tetani, 100 Hz for 500 ms at 2.5-s intervals; Fig. 1A) were elicited using an extracellular 200 mM NaCl-filled electrode placed near the fiber of interest. Green x-y images were acquired at the beginning of the electrical stimulation, and the first red image was taken immediately after completion of the green image (690 ms after initiation of electrical stimulation). Sequential green and red x-y images were taken after 1, 5, 10, 20, 30, and 40 tetani and 1, 7.5, 15, and 20 min after the last tetanus. Complete characterization of rhod 2 (Fig. 2C) and mt-pericam (Fig. 3C) fluorescence decay following five successive tetani was assessed from x-y images taken at defined times throughout the subsequent 10 min following cessation of the final tetanus.

Fig. 2.

Myoplasmic and mitochondrial Ca²⁺ transients during tetanic stimulation. A: representative confocal images of fluo 4 (top) and rhod 2 (bottom) fluorescence in FDB fibers from 4-mo-old mice before tetanic stimulation, after 1 and 5 tetani (0.2 duty cycle), and 2.5, 30, 300, and 600 s following the 5th tetanus. Scale bars, 5 μm. B: relative change (ΔF/F₀) in rhod 2 A band and fluo 4 (myoplasmic) fluorescence during and after delivery of 1 and 5 tetanic stimuli. C: normalized triadic rhod 2 fluorescence during and after delivery of 1 and 5 tetanic stimuli. Values are means ± SE; n = 17 fibers.

Fig. 4.

Ca²⁺ uptake in triadic and longitudinal mitochondria following tetanic stimulation. A: confocal images of rhod 2 fluorescence in representative FDB fibers from 4-mo-old (top) and 1-mo-old (middle and bottom) mice before stimulation, after 1, 5, 10, and 40 tetani (0.2 duty cycle), and 1, 7.5, 15, and 20 min following the last (40th) tetanus. Scale bars, 5 μm. B and C: normalized rhod 2 A band and mitochondrial rhod 2 fluorescence, respectively, during and after repetitive tetanic stimulation in FDB fibers from 4- and 1-mo-old mice. Values are means ± SE; n = 50 (4-mo-old triadic), 45 (1-mo-old triadic), and 22 [1-mo-old longitudinal (Longit)] fibers.

Fig. 5.

Mitochondrial membrane potential is unaltered during repetitive tetanic stimulation. A: representative confocal images of fluo 4 and tetramethylrhodamine ethyl ester (TMRE) fluorescence in FDB fibers from 1- and 4-mo-old mice before tetanic stimulation, after 1 and 10 tetani, and 1 and 20 min following the last (40th) tetanus. Scale bars, 5 μm. B and C: normalized myoplasmic fluo 4 fluorescence and triadic TMRE fluorescence, respectively, during and after repetitive tetanic stimulation in FDB fibers from 1- and 4-mo-old mice. Values are means ± SE; n = 5 fibers.

Fig. 3.

Transient mitochondrial Ca²⁺ uptake during twitch stimulation and sustained mitochondrial Ca²⁺ uptake following repetitive tetanic stimulation in mitochondrial-targeted ratiometric pericam (mt-pericam)-expressing fibers. A: representative confocal images of mt-pericam-expressing FDB fibers isolated from 4-mo-old mice before tetanic stimulation, after 1 and 5 tetani (0.2 duty cycle), and 2.5, 30, 300, and 600 s following the 5th and final tetanus. Scale bars, 10 μm. B: changes in mt-pericam fluorescence (402-nm excitation) monitored by confocal line scan imaging during twitch stimulations. Arrow indicates time (t = 0) of electrical stimulation. C: sustained mitochondrial Ca²⁺ increase and slow decay over 10 min following a series of 5 repetitive tetanic stimulations. Values are means ± SE; n = 6 (B) and 27 (C) fibers.

To generate a mean fluorescence value for triadic mitochondria from confocal x-y images, a 2-μm-thick line drawn along the longitudinal axis of the fiber (∼20 μm long) was used to generate a mean x-y ratio profile, with x values representing fiber length and y values representing the transversely averaged fluorescence ratio. Values for profile peaks (I band fluorescence) and troughs (A band fluorescence) following stimulation were normalized to their respective values at rest. Relative changes in triadic mitochondrial Ca²⁺ in response to stimulation were expressed as the difference between I and A band rhod 2 fluorescence within the same profile (I − A) and then normalized to the corresponding value prior to stimulation. For longitudinal clusters of mitochondria, profiles were centered over the A band.

Real-time confocal line scan imaging and analysis of mitochondrial Ca²⁺ accumulation during twitch stimulation.

Acutely dissociated, mt-pericam-expressing fibers were bathed in Ringer solution (see above) supplemented with 35 μM BTS to inhibit cell movement. A series of individual twitch transients were initially elicited using an extracellular electrode placed adjacent to the fiber of interest (1-ms pulse duration at 2 Hz). Real-time changes in relative mt-pericam fluorescence during individual twitches were monitored in confocal line scan mode (2 ms/line, 1,024 lines) across a 50-μm line drawn along the longitudinal axis of the fiber. The fiber was then stimulated with five repetitive tetani (100 Hz for 500 ms at 2.5-s intervals) followed by a series of x-y images taken over the subsequent 10 min, as described above for the rhod 2 experiments. A 402-nm laser was used to excite mt-pericam, and fluorescence emission was detected at 515 ± 30 nm. Images were acquired using a confocal microscope (Nikon Eclipse C1 Plus) equipped with a ×40 1.3 NA oil objective (SuperFluor). Real-time changes in relative mt-pericam fluorescence during twitch stimulation were assessed within a 1-μm-thick line corresponding to the mt-pericam fluorescence within the I band. Under the excitation conditions used here (402 nm), mt-pericam fluorescence decreases with an increase in Ca²⁺ binding (21). All x-y and line scan images were processed and analyzed offline using National Institutes of Health ImageJ and AutoQuant AutoDeblur and AutoVisualize software. All analyses were conducted on raw images, but for display purposes only, a two-dimensional blind deconvolution was used to minimize out-of-focus fluorescence.

Electron microscopy analysis.

FDB muscles were fixed in situ with 3.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.2) at room temperature following 1 h of exposure to isotonic or hypertonic high-Ca²⁺ Ringer solution (see below). Small portions of muscle were postfixed, embedded, stained en bloc, and sectioned for electron microscopy (EM), as described previously (4). CRUs and mitochondria, as well as the incidence of CRUs coupled to mitochondria, were marked and counted (Table 1) only in the subsarcolemmal region of fibers (1 μm deep) in nonoverlapping micrographs taken at ×17,700 magnification of longitudinal sections. Images were taken randomly for each fiber as it first appeared in the field of view as the grid was scanned, with care taken to avoid nuclear regions. If an individual mitochondrion extended from one band to the other or across the Z line, it was counted twice.

Table 1.

Quantitative analysis of mitochondria, CRUs, and CRU-mitochondrion pairing in subsarcolemmal regions of FDB muscle fibers

	Sampling	A	B	C	D
Sampling		Total Subsarcolemmal Length Analyzed, μm	No. of CRUs/100 μm	No. of Mitochondria/100 μm	No. of CRUs Associated With Mitochondria/100 μm
0.5 mo	576	57 ± 34	50 ± 42	5 ± 10 (10%)
1 mo	488	67 ± 43	50 ± 40	18 ± 21 (24%)
2 mo	486	82 ± 46	60 ± 52	34 ± 29 (40%)
4 mo	352	114 ± 46	112 ± 60	74 ± 42 (67%)

Values are means ± SD. Values in parentheses in column D represent percentage of Ca²⁺ release units (CRUs) coupled (or associated) to mitochondria. Sampling consisted of 2 mice for each time point and ≥20 flexor digitorum brevis (FDB) fibers for each mouse. Frequency of CRUs in the subsarcolemmal region increases progressively with age (column B), whereas frequency of mitochondria does not change significantly at ≤2 mo of age but doubles at 4 mo of age (column C). Percentage of CRUs associated with mitochondria increases progressively from 0.5 to ≤4 mo of age (column D). Initially (0.5–1 mo of age), mitochondria are mostly grouped in clusters under the sarcolemma. Only later are they targeted to parajunctional positions. Significant increase in frequency of CRUs and mitochondria from 2 to 4 mo of age contributes to increase in CRU-mitochondrion pairs. Statistical significance was determined by Student's t-test (P < 0.0005).

Confocal imaging and analysis of osmotic shock-induced Ca²⁺ sparks.

FDB fibers were plated on glass coverslips in isotonic Ringer solution (in mM: 140 NaCl, 4 KCl, 1 MgSO₄, 1.8 CaCl₂, 5 NaHCO₃, 10 glucose, and 10 HEPES, pH 7.4, ∼300 mosM), loaded with 10 μM fluo 4-AM for 60 min at room temperature, and then exposed for 10 min to dye-free solution. To induce Ca²⁺ sparks, fibers were immersed in hypertonic high-Ca²⁺ Ringer solution (in mM: 140 NaCl, 4 KCl, 1 MgSO₄, 50 CaCl₂, 5 NaHCO₃, 10 glucose, and 10 HEPES, pH 7.4, ∼450 mosM) (34). Ca²⁺ sparks were recorded using a confocal microscope (Nikon Eclipse C1 Plus) equipped with a ×40 1.3 NA oil objective (SuperFluor). With the focal plane centered along the fiber periphery, a series of line scans (512 pixels × 1,024 lines, 2 ms/line) parallel to the long axis of the fiber (i.e., spanning multiple sarcomeres) were acquired. Images were analyzed offline using CaSparks automated detection software. F₀ was defined as the average fluorescence level immediately before each automatically detected event. Broad event detection limits were set to identify all events with amplitudes of 0.2–6 ΔF/F, full widths at half-maximal amplitude (FWHM) of 0.5–50 μm, rise times of 0.5–100 ms, and full durations at half-maximal amplitude (FDHM) of 0.5–1,000 ms. Spark mass was calculated as amplitude ∗ 1.206 ∗ FWHM³, according to Hollingworth et al. (12), with the assumption that the scan line runs through the location of the local Ca²⁺ release event and that the three-dimensional volume of the event is isotropic in space and approximated by the product of three Gaussian functions. No correction was made for the contribution of out-of-focus events.

Statistics.

To enable comparison between different fibers within each experiment, confocal images were recorded using identical laser power and photomultiplier sensitivity and processed using identical values for contrast and brightness. Statistical significance between relative changes in myoplasmic fluo 4 fluorescence and mitochondrial rhod 2 and mt-pericam fluorescence (P < 0.05) and Ca²⁺ spark spatiotemporal properties (P < 0.001) was determined using Kruskal-Wallis one-way ANOVA with Dunn's post hoc tests.

RESULTS

Sustained mitochondrial Ca²⁺ accumulation following tetanic stimulation in rhod 2-loaded FDB fibers.

We monitored mitochondrial Ca²⁺ accumulation immediately following low- and high-frequency electrical stimulation in single mouse FDB fibers in which rhod 2 was preferentially loaded into mitochondria. In resting fibers, basal rhod 2 fluorescence was low, and no detectable change in rhod 2 fluorescence was collected in x-y confocal images 690 ms after twitch stimulation (Fig. 1C). In contrast, a robust increase in I band-delimited mitochondrial rhod 2 fluorescence (i.e., MTG-positive areas) was observed following a single high-frequency tetanus (100 Hz for 500 ms) sufficient to produce a sustained myoplasmic Ca²⁺ signal detected using mag-fluo 4 that decayed rapidly following termination of stimulation (Fig. 1, A and C). Specifically, each high-frequency tetanus evoked a rapid, robust, and sustained global myoplasmic Ca²⁺ transient (Fig. 1A, inset). The average rise time of the first twitch reached a peak within just a few milliseconds (3.86 ± 0.02 ms), and summation during the first tetanus resulted in a peak change in relative mag-fluo 4 fluorescence (ΔF/F₀ = 1.66 ± 0.24) that occurred within a few tens of milliseconds after initiation of the train (33.8 ± 1.4 ms) and remained elevated throughout the tetanus. Consistent with rapid termination of release and efficient Ca²⁺ clearance, mag-fluo 4 fluorescence abruptly decayed after termination of the tetanic stimulation train. Subsequent tetani elicited largely the same response as the first train, with a progressive reduction in peak amplitude after ∼15–20 tetani (Fig. 1B).

To more rigorously quantify the time course of the increase and decay of the mitochondrial rhod 2 signal compared with the myoplasmic Ca²⁺ transient, FDB fibers from adult mice were coloaded with myoplasmic (fluo 4) and mitochondrial (rhod 2) Ca²⁺ dyes and stimulated by a series of five successive tetani (100 Hz for 500 ms at 2.5-s intervals; Fig. 1A). The myoplasmic Ca²⁺ signal (relative change in fluo 4 fluorescence or ΔF/F₀) was maximal during the first tetanus (5.69 ± 0.63 ΔF/F₀) and decayed rapidly to near-baseline levels (0.48 ± 0.05 ΔF/F₀) after completion of the fifth tetanus (Fig. 2, A and B). As indicated above, a large and sustained increase in I band-delimited rhod 2 fluorescence occurred after the first tetanus (Fig. 2A, bottom). A minimal increase in A band-delimited rhod 2 fluorescence with kinetics identical to those of fluo 4 was presumably due to a low-level myoplasmic rhod 2 dye (Fig. 2B). Given that the rhod 2 signal within the I band region represents the sum of mitochondrial and myoplasmic components and that mitochondria are not normally present within the A band region of FDB fibers from adult mice (4), we used the difference between I band and A band rhod 2 fluorescence (I − A) as an index of the relative changes in Ca²⁺ within mitochondria located in the I band and associated with the CRU (4) (referred to here as “triadic mitochondria”). This analysis revealed a significant increase in the triadic mitochondrial Ca²⁺ signal after only one tetanus (2.72 ± 0.39) that was further increased after five successive tetani (4.12 ± 0.45). Importantly, the triadic mitochondrial Ca²⁺ signal remained significantly elevated over baseline during the subsequent 10 min, even in the absence of further stimulation (Fig. 2C). The results in Fig. 2 reveal a remarkable difference between the activation and decay kinetics of the myoplasmic and triadic mitochondrial Ca²⁺ signals following repetitive tetanic stimulation.

Mitochondrial Ca²⁺ accumulation following twitch and tetanic stimulation in mt-pericam-expressing FDB fibers.

Despite the marked difference between the time courses of the fluo 4 and triadic rhod 2 signals and the corrections made for the triadic rhod 2 signal (I − A) in Fig. 2, a potential contribution of low-level myoplasmic dye is a limitation of the rhod 2 experiments. To overcome this limitation and more rigorously evaluate changes in mitochondrial Ca²⁺ during single-twitch and repetitive high-frequency tetanic stimulation, we expressed mt-pericam in FDB muscle by in vivo electroporation. Because mt-pericam is specifically targeted to mitochondria, it enables assessment of mitochondria Ca²⁺ in the absence of any contamination from concurrent changes in myoplasmic Ca²⁺. Consistent with prior studies (25), we observed rapid and short-lived (<200 ms) mitochondrial-specific Ca²⁺ uptake during a twitch stimulation in confocal line scan measurements of mt-pericam fluorescence in expressing FDB fibers [−ΔF_{(402 nm)}/F_{0(402 nm)} = 0.19 ± 0.02; Fig. 3B]. On the other hand, five successive high-frequency tetani (100 Hz for 500 ms at 2.5 s intervals) elicited a larger (0.25 ± 0.01) and more sustained increase in mitochondrial Ca²⁺, taking >10 min to fully decay to baseline (Fig. 3, A and C). These latter results are essentially identical to those observed for rhod 2-loaded fibers in Fig. 2, confirming a robust and sustained Ca²⁺ increase in triadic mitochondria following only several brief tetani.

Sustained Ca²⁺ accumulation in triadic and longitudinal mitochondria following repetitive tetanic stimulation.

FDB fibers from young (1-mo-old) mice exhibit triadic mitochondria and longitudinal clusters of subsarcolemmal and intermyofibrillar mitochondria (4). To directly assess mitochondrial Ca²⁺ uptake in triadic and longitudinal mitochondria, we compared the magnitude and time course of changes in mitochondrial rhod 2 fluorescence following repetitive tetanic stimulation (40 tetani, 100 Hz, 500 ms, 2.5-s intervals) in FDB fibers from young (1-mo-old) and adult (4-mo-old) mice. Consistent with results shown in Figs. 1 and 2, robust triadic mitochondrial Ca²⁺ accumulation was observed in FDB fibers from 4-mo-old mice following the first tetanus, was increased for the next 5–10 tetani, and remained elevated even after 40 tetani (3.78 ± 0.42, 5.36 ± 0.52, 6.32 ± 0.54, and 6.62 ± 0.55, respectively; Fig. 4, A and C). Only a minimal A band-associated rhod 2 signal was observed in FDB fibers from young (1-mo-old) and adult (4-mo-old) mice (Fig. 4B).

Unexpectedly, the time course of mitochondrial Ca²⁺ uptake and decay was similar for CRU-paired triadic and longitudinal mitochondria in fibers from young mice (Fig. 4, A and C). The prominent triadic and longitudinal mitochondrial Ca²⁺ signals remained elevated throughout the repetitive tetanic stimulation protocol, persisted to about the same level for >1 min following the 40th tetanus, exhibited half times of decay of ∼7 min, and required 20 min to return completely to baseline levels (Fig. 4, A and C). The extremely slow decay of the triadic and longitudinal mitochondrial Ca²⁺ signals is in direct contrast to the rapid decay of the myoplasmic Ca²⁺ signal following the termination of tetanic stimulation. Thus, repetitive tetanic stimulation triggers a similar sustained accumulation of Ca²⁺ in the triadic and longitudinal mitochondrial populations.

Mitochondrial Ca²⁺ accumulation occurs in the absence of mitochondrial depolarization.

To assess potential effects of mitochondrial Ca²⁺ uptake on the mitochondrial membrane potential, we simultaneously monitored myoplasmic fluo 4 fluorescence and mitochondrial TMRE fluorescence during repetitive tetanic stimulation (Fig. 5). Similar to MTG fluorescence intensity profiles (Fig. 1C), FDB fibers loaded with TMRE exhibit clear double rows of transverse red fluorescence, consistent with efficient loading of the dye into I band-delimited triadic mitochondria. The repetitive tetanic stimulation protocol used in Figs. 1–4 to trigger robust and sustained mitochondrial Ca²⁺ uptake failed to significantly alter steady-state triadic TMRE fluorescence in FDB fibers from young (1-mo-old) and adult (4-mo-old) mice (Fig. 5, A and C). Thus, mitochondrial Ca²⁺ uptake during repetitive tetanic stimulation does not result in a sustained depolarization of the mitochondrial membrane potential. However, because the single x-y TMRE images shown in Fig. 5A were taken between each tetanus, transient changes in mitochondrial membrane potential that may have occurred during tetanic stimulation would have escaped detection. Nevertheless, the maintenance of robust electrically evoked SR Ca²⁺ release (Fig. 5B) and steady-state triadic mitochondrial TMRE fluorescence (Fig. 5C) throughout the repetitive tetanic stimulation protocol demonstrates that sarcolemmal and mitochondrial membrane potentials remained intact throughout these experiments.

Triadic and longitudinal mitochondria are continuous and structurally tethered to the SR.

We previously demonstrated that mitochondrial disposition changes during postnatal maturation of fast-twitch skeletal fibers from mice (4). Initially (at 0.5 mo of age), most mitochondria are clustered longitudinally between myofibrils or under the sarcolemma. Numbers of mitochondria and CRUs increase substantially over the subsequent few months (1–4 mo). During this time, mitochondria progressively move to the I band adjacent to CRUs (triadic mitochondria), forming two distinct transversal networks, one on either side of the Z line (Fig. 6A, open arrows). Longitudinal rows of intermyofibrillar mitochondria, frequent at 0.5–1 mo of age, are still present occasionally in fast-twitch fibers even at 2–4 mo of age (Fig. 6A, white arrows). A more detailed EM characterization of the longitudinal mitochondrial population enabled determination of morphological features that may explain the functional data shown in Fig. 4. First, longitudinal mitochondria are structurally continuous with triadic mitochondria (Fig. 6B, arrowheads). Second, triadic and longitudinal mitochondria are closely associated with the SR: triadic mitochondria are in close proximity to CRUs at the I band (Fig. 6, B and C), while longitudinal mitochondria are surrounded by nonjunctional SR (Fig. 6, B and D) at the A band (Fig. 6B). Importantly, high-magnification EM analysis revealed that triadic and longitudinal mitochondria are structurally tethered to the SR by small electron-dense strands that appear to hold the two organelles together, creating a restricted ∼10- to 15-nm-wide subcellular space (Fig. 6, C and D, black arrows).

Fig. 6.

Triadic and longitudinal mitochondria are continuous and structurally tethered to the sarcoplasmic reticulum (SR). A: representative electron micrograph (longitudinal section) of an FDB fiber from a 4-mo-old mouse highlighting location of longitudinal (white arrows) and triadic (open arrows) mitochondria. B: representative electron micrograph highlighting 1) continuity (arrowheads) between triadic (open arrows) and longitudinal (white arrow) mitochondria and 2) association with the longitudinal SR (long SR, small black arrows). C and D: under higher magnification, small electron-dense strands (insets, black arrows) tether mitochondria to the triad or the longitudinal (long) SR. Scale bars, 1 μm (A), 0.5 μm (B), and 0.25 μm (C and D).

Osmotic shock induces changes in subsarcolemmal structure.

Exposure to hyperosmotic solution elicits peripherally located local Ca²⁺ release events (or “Ca²⁺ sparks”) in skeletal muscle fibers (34). Here, we determined the effect of osmotic shock on muscle fiber ultrastructure. This treatment did not grossly alter triadic mitochondrial localization, since the characteristic double-row banded pattern of MitoTracker Red CMXRos (MTR) fluorescence was retained in FDB fibers from 4-mo-old mice following exposure to hypertonic high-Ca²⁺ Ringer solution (Fig. 7A). More detailed EM analyses confirmed the absence of major ultrastructural changes within the fiber interior following osmotic shock in fibers from 0.5- or 4-mo-old mice. On the other hand, osmotic shock induced detectable changes within subsarcolemmal regions of the fiber perimeter (i.e., within 1 μm under the surface membrane in EM images; Fig. 7, B and C). Specifically, exposure to hypertonic high-Ca²⁺ Ringer solution induced swelling of T-tubules and caveolae just beneath the surface membrane, the extent of which was variable within a given field of view. Importantly, osmotic shock produced very similar structural changes in FDB fibers from 0.5- and 4-mo-old mice. The only significant difference between the two postnatal time points was a higher percentage of fibers from young (0.5-mo-old) mice exhibiting subsarcolemmal and intermyofibrillar clusters of mitochondria, consistent with the progressive relocation of mitochondria from longitudinal rows to parajunctional positions within the I band (Table 1) during postnatal development, as described previously (4).

Fig. 7.

Exposure to hypertonic high-Ca²⁺ osmotic shock induces swelling of T-tubules and caveolae. A: representative confocal images of MitoTracker Red CMXRos (MTR) fluorescence in a FDB fiber from a 4-mo-old mouse before (left) and after (right) exposure to hypertonic high-Ca²⁺ osmotic shock. B and C: electron micrographs (longitudinal sections) of FDB fibers from 0.5- and 4-mo-old mice exposed to isotonic or hypertonic high-Ca²⁺ solution before fixation. Left: subsarcolemmal regions; right: T-tubule profiles within triads at higher magnification. Small black arrows, caveolae; large black arrows, dilated caveolae; small white arrows, T-tubules; large white arrows, dilated T-tubules.

Altered frequency and spatiotemporal properties of osmotic shock-induced Ca²⁺ sparks during postnatal maturation.

We compared the incidence and properties of Ca²⁺ sparks induced by exposure to hypertonic high-Ca²⁺ (450 mosM) Ringer solution at different stages of postnatal maturation (0.5, 1, 2, and 4 mo) that exhibit marked differences in mitochondrion-CRU tethering (4). After exposure to hypertonic high-Ca²⁺ solution, FDB fibers of all ages exhibited local Ca²⁺ release events (Fig. 8A). Activity persisted for the subsequent hour of experimentation, primarily at the fiber periphery. Indeed, triads potentially involved in mediating peripherally localized Ca²⁺ sparks were found within a few micrometers of the sarcolemma (Fig. 7, B and C, Table 1). Because the external solution contained Ca²⁺, a contribution of Ca²⁺ influx cannot be excluded. However, the quantal nature of the events argues against nonspecific membrane tears.

Fig. 8.

Properties of osmotic shock-induced Ca²⁺ sparks in FDB fibers during postnatal development. A: representative x-t line scan images of Ca²⁺ sparks in FDB fibers from 0.5-, 1-, 2-, and 4-mo-old mice. For display purposes, images were processed using a Gaussian smoothing function and median filter and then normalized to background. Arrows mark representative “Ca²⁺ sparks” and “Ca²⁺ bursts.” B: peak Ca²⁺ spark amplitude. C: full width at half-maximal amplitude (FWHM). D: Ca²⁺ spark mass calculated as amplitude ∗ 1.206 ∗ FWHM³ (12). E and F: raw Ca²⁺ spark frequency and Ca²⁺ spark frequency normalized to Ca²⁺ release unit (CRU) density, with 21 ± 5, 43 ± 9, 69 ± 12, and 88 ± 25 CRUs/100 μm² used for fibers from 0.5-, 1-, 2-, and 4-mo-old mice, respectively (4). G: correlation between normalized Ca²⁺ spark frequency and tether density, with 0.8, 4.0, 10.6, and 15.6 tethers/100 μm² used for fibers from 0.5-, 1-, 2-, and 4-mo-old mice, respectively. Values are means ± SE; n = 875 (0.5-mo-old), 710 (1-mo-old), 1,286 (2-mo-old), and 2,278 (4-mo-old) events. *P < 0.001 vs. 4-mo-old mice.

Ca²⁺ spark frequency and morphology varied throughout postnatal maturation. At all ages, brief (∼20 to 30 ms) local Ca²⁺ release events (termed Ca²⁺ sparks) with spatiotemporal properties similar to evoked Ca²⁺ sparks identified by others in mammalian skeletal muscle preparations (6, 13, 14, 34) were observed (Fig. 8A). In addition, longer-duration (>100 ms) events (Fig. 8A), analogous to previously termed Ca²⁺ bursts (34), were also detected. However, since the spatiotemporal properties of the short- and long-duration events exhibited similar changes with age, all subsequent analyses were conducted on the entire population of events and, hereafter, are collectively referred to as Ca²⁺ sparks.

Mean peak Ca²⁺ spark amplitude, spatial width, and mass at the different postnatal ages (0.5, 1, 2, and 4 mo) are summarized in Fig. 8, B–D. Mean peak Ca²⁺ spark amplitude was significantly lower in fibers from 0.5-mo-old mice (0.3 ± 0.01 ΔF/F) than in fibers from 1-, 2-, and 4-mo-old mice, which exhibited similar average peak amplitudes (0.4 ± 0.01, 0.4 ± 0.01, and 0.4 ± 0.01 ΔF/F, respectively; Fig. 8B). A similar postnatal trend was also observed for average Ca²⁺ spark rise time (data not shown). In contrast, average FWHM peaked at 1 mo (38.2 ± 1.0 ms) and was reduced at 0.5, 2, and 4 mo (29.5 ± 0.8, 33.3 ± 0.7, and 33.9 ± 0.5 ms, respectively; Fig. 8C). Calculated spark mass [amplitude ∗ 1.206 ∗ FWHM³ (12)] was greater at 1 mo than at all other time points (1.8 ± 0.2, 2.8 ± 0.2, 2.1 ± 0.1, and 1.8 ± 0.1 μm³ for 0.5, 1, 2, and 4 mo, respectively; Fig. 8D).

The most dramatic effect of postnatal maturation was on Ca²⁺ spark frequency (Fig. 8, E–G). Raw Ca²⁺ spark frequency followed the identical pattern of amplitude and FDHM (data not shown), being lowest at 0.5 mo (76.1 ± 3.7 events/min) and significantly higher in FDB fibers from 1-, 2- and 4-mo-old mice (115.0 ± 9.1, 110.1 ± 5.7, and 107.8 ± 4.3 events/min, respectively; Fig. 8E). This trend is consistent with the increase in the CRU number and density during postnatal maturation in subsarcolemmal (i.e., <1 μm from the sarcolemma; Table 1) and internal regions of myofibers (4). To evaluate Ca²⁺ spark frequency per CRU, we normalized raw Ca²⁺ spark frequencies to the CRU density at each developmental stage. Osmotic shock-induced Ca²⁺ sparks in our experiments occurred primarily at the fiber periphery, within 5 μm from the surface membrane. Given this relatively large spatial range from the surface membrane, we normalized the raw Ca²⁺ spark frequencies using the corresponding “internal” CRU densities at each developmental stage (4). Significantly, Ca²⁺ spark frequency per CRU density decreased progressively throughout postnatal development (3.6 ± 0.2, 2.7 ± 0.2, 1.6 ± 0.1, and 1.2 ± 0.1 events/min/CRU/100 μm² at 0.5, 1, 2, and 4 mo, respectively; Fig. 8F). More importantly, a strong linear correlation (r² = 0.95; Fig. 8G) was observed between the normalized Ca²⁺ spark frequency and the mitochondrion-CRU tether density determined previously (4).

DISCUSSION

This study provides important new information with regard to SR-mitochondrial communication in skeletal muscle. 1) Mitochondrial Ca²⁺ uptake was robust and long-lived (lasting several minutes) following repetitive tetanic stimulation (Figs. 2–4) in the absence of sustained depolarization of the mitochondrial membrane potential. 2) Unexpectedly, sustained mitochondrial Ca²⁺ uptake following repetitive tetanic stimulation was similar for triadic and longitudinal mitochondrial populations (Fig. 4), even though the two different populations are located at substantially different positions relative to the CRU. 3) Sustained mitochondrial Ca²⁺ uptake was not accompanied by a steady-state change in mitochondrial membrane potential (Fig. 5). 4) As observed previously for CRU-coupled mitochondria (4), high-resolution EM identified restricted sites of contact between longitudinal mitochondria and the nonjunctional SR that are mediated by electron-dense tethers (Fig. 6). 5) A progressive increase in retrograde suppression of local SR Ca²⁺ release during postnatal maturation occurs in direct correspondence with a parallel increase in mitochondrial triadic positioning (Fig. 8). These findings indicate that mitochondrial tethering to the CRU is more critical for Ca²⁺ spark suppression than for sustained mitochondrial Ca²⁺ uptake during tetanic stimulation. The relationship of these findings to prior published studies and their implications for skeletal muscle function are discussed in detail below.

Implications of sustained mitochondrial Ca²⁺ accumulation following tetanic stimulation.

Consistent with prior studies (25), mitochondrial Ca²⁺ uptake was transient (<200 ms) following a single electrically evoked twitch (Fig. 3B). On the other hand, mitochondria in FDB fibers from young (1-mo-old) and adult (4-mo-old) mice exhibited robust and sustained Ca²⁺ uptake following repetitive tetanic stimulation (Figs. 2–4). Importantly, we confirmed sustained mitochondrial Ca²⁺ uptake following repetitive tetanic stimulation in rhod 2-loaded (Figs. 2 and 4) and mt-pericam-expressing (Fig. 3) fibers. In the absence of calibration of these signals, the absolute level of the mitochondrial Ca²⁺ increase during tetanic stimulation remains unknown. However, the actual increase in mitochondrial free Ca²⁺ is likely to be rather modest, given the lack of a detectable depolarization of the mitochondrial membrane potential, a characteristic of mitochondrial Ca²⁺ overload, or overt muscle fiber damage following the prolonged repetitive tetanic stimulation used in this study.

Our results are in contrast to those of Lannergren et al. (18), who reported negligible mitochondrial Ca²⁺ uptake in toe fibers following tetanic stimulation. There are several possible explanations for this difference. Unlike the study of Lannergren et al., our experiments were conducted in the presence of BTS to block contraction (8, 28) and minimize movement artifacts. Given that BTS inhibits myosin ATPase activity (28), total ATP consumption during tetanic stimulation would be reduced in our experiments. As a consequence, there is a reduced need for mitochondrial ATP production and subsequent dissipation of the mitochondrial membrane potential, thus potentially enhancing the driving force for Ca²⁺ uptake. Alternatively, BTS may increase Ca²⁺ availability for mitochondrial Ca²⁺ uptake by reducing crossbridge cycling and Ca²⁺ binding to troponin C.

On the other hand, our results are consistent with those of Bruton et al. (7), who demonstrated significant mitochondrial Ca²⁺ accumulation during tetanic stimulation of mouse extensor digitorum longus and soleus muscle fibers, and Shkryl et al. (29), who reported mitochondrial Ca²⁺ uptake during caffeine-induced Ca²⁺ release in permeabilized extensor digitorum longus and soleus fibers. Furthermore, Aydin and colleagues (1) reported significant mitochondrial Ca²⁺ uptake in myopathic FDB fibers from Tfam (mitochondrial transcription factor A) knockout mice, although only a slight increase was observed in control fibers after 25 tetani. The smaller increase in mitochondrial Ca²⁺ uptake in control fibers observed by Aydin et al. could be due to the three- to four-times-longer duration of dye loading used in the prior study, resulting in significantly elevated resting rhod 2 fluorescence levels and, thus, a smaller relative increase in rhod 2 fluorescence upon stimulation. In any event, our results are the first to demonstrate significant mitochondrial Ca²⁺ accumulation in intact wild-type FDB fibers using a physiological tetanic stimulation paradigm.

The combination of efficient mitochondrial Ca²⁺ uptake and slow Ca²⁺ removal results in mitochondria functioning as “temporal Ca²⁺ integrators” during tetanic stimulation. Consequently, mitochondrial Ca²⁺ accumulation is sustained following repetitive tetanic stimulation and, thus, ideally suited to promote prolonged Ca²⁺ activation of ATP generation (20, 31) precisely when it is most needed, i.e., under conditions of intense muscle activity.

Potential mechanisms for sustained Ca²⁺ accumulation into triadic and longitudinal mitochondria following tetanic stimulation.

Since mitochondria in the I band are tethered to CRUs (4), we initially expected Ca²⁺ uptake to be greater in triadic than longitudinal mitochondria. Thus, our demonstration of similar mitochondrial Ca²⁺ uptake for both mitochondrial populations was unexpected. What mechanisms could account for comparable activity-dependent mitochondrial Ca²⁺ uptake in triadic and longitudinal mitochondrial populations? As one possibility, longitudinal and triadic mitochondria may respond to global Ca²⁺ signals, rather than restricted Ca²⁺ release microdomains within the immediate vicinity of RYRs located in the CRU. Indeed, triadic mitochondria are positioned on the side of the CRU opposite to the site of Ca²⁺ release, such that the minimal distance between the site of release and the closest outer mitochondrial membrane is usually ∼130 nm (4). For the few triadic mitochondria that run along the side of triads and extend into the longitudinal population (Fig. 6B), this distance may be somewhat reduced. Simulations of Ca²⁺ release in an isotropic medium indicate that Ca²⁺ release microdomains dissipate markedly only a few tens of nanometers and almost entirely ∼100 nm away from the point of release (30). Thus, mitochondria will experience a significant Ca²⁺ release microdomain only in cases where the distance between RYRs and the mitochondrial membrane is significantly <100 nm. Such cases are extremely rare in adult fibers but somewhat more frequent in developing muscle.

Given the low affinity of mitochondrial Ca²⁺ uptake mechanisms and location of mitochondria outside the RYR Ca²⁺ release microdomain, Ca²⁺ uptake by triadic and longitudinal mitochondria may involve a unique form of SR-mitochondrial communication. For example, continuity is observed between triadic and longitudinal mitochondrial matrixes, indicating that the two populations are not structurally separated. Thus, Ca²⁺ uptake by one population may influence the other via intramatrix transfer. In addition to a direct continuity, the SR is connected to triadic and longitudinal mitochondria by small strands or tethers that hold the two organelles together. Tethers create a restricted ∼10- to 15-nm-wide interorganelle space, referred to here as “SR-mitochondrial nanodomains” (Fig. 9). Although RYR Ca²⁺ release channels are unlikely to be present within these nanodomains, SERCA expression, which is extremely high throughout the nonjunctional SR (9, 15), should be high. Thus, mitochondrial Ca²⁺ uptake following tetanic stimulation could, in part, involve Ca²⁺ accumulation within SR-mitochondrial nanodomains during SERCA unidirectional reverse flux (or backflux) (27a, 27b). SERCA Ca²⁺ backflux, which is enhanced by ADP, may be sufficient (following repetitive tetanic stimulation) to limit or even surpass SR Ca²⁺ uptake and, thus, allow for accumulation of Ca²⁺ on the myoplasmic side of the SR membrane. By virtue of being tethered to nonjunctional SR (4), triadic and longitudinal mitochondria could experience a similar SERCA backflux during repetitive tetanic stimulation sufficient to activate mitochondrial Ca²⁺ uptake. Regardless of the mechanism, our observation that Ca²⁺ uptake following tetanic stimulation is similar for triadic and longitudinal mitochondria indicates that targeting to the CRU is not required for sustained mitochondrial Ca²⁺ uptake.

Fig. 9.

Schematic representation of intimate interactions between SR and both triadic and longitudinal mitochondria. Triadic and longitudinal mitochondria are associated with the SR by small strands or tethers, which hold the membranes of the 2 organelles close together, creating a restricted subcellular space ∼10–15 nm wide, referred to as the “SR-mitochondrial nanodomain.” Yellow, SR; white, T-tubules; blue, ryanodine receptor 1 feet; green, mitochondria; gray, calsequestrin; red, SR-mitochondrial tethers; cyan, nanodomains.

Increased mitochondrion-CRU coupling correlates with Ca²⁺ spark suppression.

Our results suggest that mitochondrion-CRU targeting plays a more important role in retrograde suppression of local Ca²⁺ release. Retrograde suppression of local Ca²⁺ sparks in mammalian skeletal muscle (13, 14, 19) is facilitated by close proximity of mitochondria to CRUs, although mitochondria-independent mechanisms of Ca²⁺ spark suppression may also be at play (16). In support of the former, we found that the frequency of osmotic shock-induced Ca²⁺ sparks per CRU density decreased threefold in FDB fibers during postnatal maturation, in direct correspondence with a parallel increase in mitochondrion-CRU pairing and tether density. Although this reduction in Ca²⁺ spark frequency could also reflect other changes in the CRU that occur during development, these findings are consistent with the local redox control mechanism for mitochondrial-mediated inhibition of local SR Ca²⁺ release championed by Shirokova and colleagues (13, 14, 19).

How might osmotic shock preferentially overcome Ca²⁺ spark suppression on the fiber periphery? Swelling of caveolae and T-tubules at the periphery may disrupt dihydropyridine receptor inhibition of spontaneous RyR1 Ca²⁺ release. In addition, changes in subsarcolemmal structure induced by osmotic shock may interfere with proper redox signaling within the CRU. Specifically, we found that osmotic shock produced dramatic structural changes in caveolae, small (50- to 100-nm) Ω-shaped invaginations of the sarcolemma (26). These structural changes in caveolae may disrupt scaffolding of critical signal transduction elements (e.g., caveolin-3 and neuronal nitric oxide synthase) (11, 26, 33) that enhance local ROS/RNS stress. Tethering of mitochondria to the CRU would ensure that mitochondrial ROS/RNS detoxification systems are strategically positioned to minimize this insult, although potentially less so at the fiber periphery, where caveolae are located. Future work is needed to test the validity of this hypothesis and to determine if the strong correlation between Ca²⁺ spark frequency and mitochondrial CRU tethering also holds for other triggers of Ca²⁺ sparks in skeletal muscle (e.g., strenuous exercise).

Conclusions and future perspectives.

In summary, we provide new insights into bidirectional SR-mitochondrial communication in skeletal muscle. Sustained mitochondrial Ca²⁺ uptake following tetanic stimulation is not restricted to triadic mitochondria and does not result in sustained mitochondrial depolarization. Conversely, a strong correlation exists between the progressive increase in mitochondrion-CRU tethering during postnatal maturation and a parallel reduction in susceptibility to Ca²⁺ spark activation, consistent with Ca²⁺ spark suppression being mediated by a retrograde signal from energized mitochondria to the adjacent CRU (13, 14, 19). Thus, mitochondrial tethering to the CRU is more critical for Ca²⁺ spark suppression than for sustained mitochondrial Ca²⁺ uptake. The results provide evidence for differential bidirectional SR-mitochondrial Ca²⁺ cross talk in skeletal muscle and justify future investigations to determine the role of this signaling mechanism in exercise, fatigue, and muscle disease. In addition, the importance of bidirectional SR-mitochondrial signaling may differ between fast- and slow-twitch muscle fibers, as each differs with respect to mitochondrial content and positioning, relative oxidative vs. glycolytic activities, and ROS production/scavenging capacities.

GRANTS

This research was supported by National Institutes of Health Grants AR-044657 (to R. T. Dirksen) and T32 DE-07202 (to A. E. Rossi), a Telethon Research Grant (GGP08153 to F. Protasi), and the Academia Dei Lincea Fund (to L. Wei).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.