Absence of physiological Ca2+ transients is an initial trigger for mitochondrial dysfunction in skeletal muscle following denervation
© The Author(s). 2017
Received: 8 December 2016
Accepted: 8 March 2017
Published: 10 April 2017
Motor neurons control muscle contraction by initiating action potentials in muscle. Denervation of muscle from motor neurons leads to muscle atrophy, which is linked to mitochondrial dysfunction. It is known that denervation promotes mitochondrial reactive oxygen species (ROS) production in muscle, whereas the initial cause of mitochondrial ROS production in denervated muscle remains elusive. Since denervation isolates muscle from motor neurons and deprives it from any electric stimulation, no action potentials are initiated, and therefore, no physiological Ca2+ transients are generated inside denervated muscle fibers. We tested whether loss of physiological Ca2+ transients is an initial cause leading to mitochondrial dysfunction in denervated skeletal muscle.
A transgenic mouse model expressing a mitochondrial targeted biosensor (mt-cpYFP) allowed a real-time measurement of the ROS-related mitochondrial metabolic function following denervation, termed “mitoflash.” Using live cell imaging, electrophysiological, pharmacological, and biochemical studies, we examined a potential molecular mechanism that initiates ROS-related mitochondrial dysfunction following denervation.
We found that muscle fibers showed a fourfold increase in mitoflash activity 24 h after denervation. The denervation-induced mitoflash activity was likely associated with an increased activity of mitochondrial permeability transition pore (mPTP), as the mitoflash activity was attenuated by application of cyclosporine A. Electrical stimulation rapidly reduced mitoflash activity in both sham and denervated muscle fibers. We further demonstrated that the Ca2+ level inside mitochondria follows the time course of the cytosolic Ca2+ transient and that inhibition of mitochondrial Ca2+ uptake by Ru360 blocks the effect of electric stimulation on mitoflash activity.
The loss of cytosolic Ca2+ transients due to denervation results in the downstream absence of mitochondrial Ca2+ uptake. Our studies suggest that this could be an initial trigger for enhanced mPTP-related mitochondrial ROS generation in skeletal muscle.
KeywordsE-C coupling Calcium imaging Calcium signaling Calcium intracellular release Denervation Mitochondria
Skeletal muscle is responsible for voluntary movements of the entire body. Because it comprises around 40% of whole-body lean mass of a human, skeletal muscle is also essential for maintaining the homeostasis of the whole-body metabolism . Skeletal muscle contraction is under the control of motor neurons. In some disease states, the interaction between motor neuron and skeletal muscle is lost, leading to paralysis and muscle atrophy . Muscle atrophy is defined as a decrease in the muscle cell size and the imbalance between protein synthesis and degradation. The molecular mechanisms underlying protein metabolism in muscle atrophy have been extensively evaluated [3–8]. It is believed that skeletal muscle atrophy is caused by the disturbance of signaling networks, in which mitochondria may play a major role. In fact, mitochondria occupy about 10–15% of the muscle fiber volume . They are not only essential for energy supply but also determine the survival or death of muscle fibers. It has been shown that denervation of skeletal muscle induces a dramatic increase in mitochondrial ROS production . However, the initial cause of the mitochondrial ROS production in denervated skeletal muscle remains elusive .
Muscle cells use Ca2+ as a messenger to control events ranging from activation of contraction to cell death. Defective intracellular Ca2+ signaling has been linked to skeletal muscle dysfunction during aging [12, 13] and in muscular dystrophy (mdx) [14–17]. In non-muscle cells, mitochondria dynamically transport Ca2+ and modify its flux into the endoplasmic reticulum, nucleus, and across the plasma membrane to such an extent that they have been named “the hub of cellular Ca2+ signaling” . There is strong evidence that mitochondria may have a similar role in skeletal muscle. We previously demonstrated that mitochondria take up Ca2+ during excitation-contraction (E-C) coupling following rapid calcium transients in skeletal muscle . We also established that malfunction of this mechanism contributes to neuromuscular degeneration in amyotrophic lateral sclerosis . Mitochondrial Ca2+ uptake is believed to help regulate mitochondrial metabolism and ATP synthesis, so that the energy demands of muscle contraction are met . However, Ca2+ overload in mitochondria is also a pathological stimulus of ROS generation . It has been shown that prolonged muscle denervation leads to an increased resting cytosolic free Ca2+ level, that in turn overloads mitochondria, stimulating ROS production [23, 24]. Following denervation, no action potential can be initiated; therefore, physiological Ca2+ transient is lost in the denervated muscle fibers. One essential question to ask is how mitochondria respond to the cessation of physiological Ca2+ transients. While published studies mainly focused on the effect of a steady state level of intracellular Ca2+ on mitochondrial function, it is not known whether the dynamic change of Ca2+ level inside mitochondria in response to the intracellular Ca2+ transients is also a player in regulating mitochondrial function.
In this study, we used a transgenic mouse model carrying a mitochondrial biosensor mt-cpYFP, which produces mitoflash signal as a functional indication of ROS-related mitochondrial metabolic function [25, 26]. This model allows us to detect the early changes of ROS-related mitochondrial metabolic function in live skeletal muscle in response to the denervation process. Under voltage-clamp condition, we found that the loss of physiological Ca2+ transients or mitochondrial Ca2+ uptake could be an initial trigger for mitochondrial dysfunction with increased mitochondrial ROS production in skeletal muscle fibers following denervation.
Generation of transgenic mice
The plasmid (mt-cpYFP/pUCCAGGS) used to generate this transgenic line (mt-cpYFP) is the same as the one developed by Dr. Heping Cheng’s Laboratory to generate the original mt-cpYFP transgenic mice [26, 27]. The genetic background for this transgenic mouse model is B6SJL from the Jackson Laboratory. The mt-cpYFP mice at the age of 2.5–3 months were used. All experiments were carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Protocols on usage of mice were approved by the Institutional Animal Care and Use Committee of Rush University, University of Missouri at Kansas City and Kansas City University of Medicine and Bioscience.
Muscle denervation procedure
Muscle denervation was performed by transection of the sciatic nerve. During a denervation procedure, the mouse was anesthetized with constant-flow isoflurane inhalation and a small incision was made in the mid-posterolateral area of the thigh, and the sciatic nerve was isolated. In one hind limb, the sciatic nerve was severed and a ~5 mm section was removed. The ends of the nerve were sutured to prevent nerve regrowth. For control experiments (sham), the sciatic nerve was exposed in the contralateral hind limb without being severed. The incisions in both legs were closed again with silk sutures, and the animal was euthanized after 24 h for experiments.
Isolation of FDB fibers
The animals were euthanized by CO2 inhalation followed by cervical dislocation, and the flexor digitorum brevis (FDB) muscles were removed for imaging studies. Individual FDB muscle fibers were isolated using a modified collagenase-digestion method described previously . Briefly, FDB muscles were digested in modified Krebs solution (0 Ca2+) containing 0.2% type I collagenase (Sigma), for 35 min at 37 °C. After digestion, muscles were kept in collagenase-free Krebs solution (with Ca2+ and 10 mM glucose) at 4 °C and used for imaging within 6 h. All experiments were conducted under the same isolation procedure and the same time window. In addition, both Sham and Denervation procedures were conducted in the same mouse, thus the paired sham and denervated FDB muscles were isolated from the same mouse for each experiment.
Confocal imaging of mitoflash (cpYFP) signals and image analysis
Zeiss LSM 510 live confocal microscope was used for imaging cpYFP fluorescence. Images were captured with a ×40, 1.2 NA water immersion objective at a sampling rate of 1 s/frame. Dual excitation of mt-cpYFP was achieved by alternating excitation at 405 and 488 nm, and the emission was collected at >505 nm. Experiments were performed at room temperature (23 °C). Time-lapse confocal images were analyzed using custom-developed algorithms written in Interactive Data language (IDL)  and ImageJ (NIH). Motion artifacts, background subtraction and photo-bleach correction were taken into account before automatic flash detection, using the built in custom-software.
FDB fibers isolated from denervated and sham-operated mt-cpYFP mice were bathed in Tyrode’s solution. Individual fibers were electrically stimulated with a stimulation protocol described previously with some modifications . Briefly, extracellular platinum wire electrodes were placed in parallel to the cell of interest. A single field stimulation of 350 ms in duration, consisting of 0.5 ms (8–12 V) pulses applied at a frequency of 40 Hz was generated. Fibers that did not visually respond to stimulation were excluded from the analysis. Mitoflash signal was monitored by collecting consecutive x−y time series confocal images (100 images) immediately before electrical stimulation, 10 and 120 s after termination of the field stimulation.
Electroporation and gene expression in FDB muscle of adult mice
The procedure was modified from our previous study [20, 30]. The anesthetized mice were injected with 10 μl of 2 mg/ml hyaluronidase dissolved in sterile saline at the ventral side of the hind paws through a 29-gauge needle. One hour later, 5–10 μg of plasmid DNA of pCDNA3/mt11-YC3.6 in 10 μl of sterile saline were injected into the same sites. Fifteen minutes later, two electrodes (gold-plated stainless steel acupuncture needles) were placed at the starting lines of paw and toes, separated by about 9 mm. Twenty pulses of 100 V/cm with 20 ms duration were applied at 1 Hz (ECM 830 Electro Square Porator, BTX Harvard Apparatus). Seven days later, the mice were euthanized and FDB muscles were removed for functional studies.
Voltage clamp of FDB muscle fibers
The method was modified from our previous study [20, 31]. Muscle fibers expressing mt11-YC3.6 were patched with 0.6–1.0 MΩ pipettes filled with a cesium glutamate-based solution containing 120 mM cesium glutamate, 5 mM EGTA, 10 mM glucose, 10 mM Tris base, 5 mM ATP, 10 mM phosphocreatine, 1 mM Mg2+, 100 nM Ca2+, and 50 μM M x-rhod-1. An Axopatch 200B amplifier (Axon Instruments, Foster City, CA) was used for whole-cell patch clamp. The fiber was clamped at −80 mV, and changes in cytosolic Ca2+ transients were recorded following application of depolarizing voltages. The external solution contained 140 mM triethanolamine CH3SO3H, 10 mM Hepes, 1 mM CaCl2, 3.5 mM MgCl2, 1 mM 4-aminopyridine, 0.3 mM LaCl3, 0.5 mM CdCl2, 0.5 μM tetrodotoxin, and 50 μM N-benzyl-p-toluenesulfonamide.
Confocal imaging of YC3.6, x-rhod-1 and MitoSOX Red in FDB muscle fibers
We have developed a method to simultaneously record the images of YC3.6 for mitochondrial Ca2+ signaling and x-rhod-1 for cytosolic Ca2+ signaling . Separate excitation and emission wavelengths were applied for simultaneous recording of x-rhod-1 and YC3.6 signals. x-rhod-1 was excited at 594 nm, and its fluorescence was collected at 600–680 nm. mt11-YC3.6 was excited at 458 nm, and its fluorescence f 1 was collected at 470–520 nm and f 2 at 520–580 nm. MitoSOX Red (M36008, Invitrogen) was used to evaluate mitochondrial superoxide production level. FDB muscle fibers were incubated with 0.5 μM MitoSOX Red in our modified Krebs solution at 37 °C for 10 min. MitoSOX Red was excited at 514 nm, and its fluorescent images were collected at 570–630 nm under the Leica SP8 confocal microscope.
Isolation of mitochondrial fraction from skeletal muscles
Crude mitochondrial fractionations from skeletal muscle were first obtained using a method previously reported [32, 33] with some modifications. Briefly, fresh skeletal muscles were removed from the mice and placed in ice-cold phosphate-buffered saline (PBS) with 10 mM EDTA. The muscles were suspended in 10 ml/gram weight ice-cold homogenization buffer (100 mM sucrose, 10 mM EDTA, 100 mM Tris-HCl, 46 mM KCl, pH 7.4 with 5 mg/ml BSA and proteinase inhibitor (Thermo Fisher)), minced into small pieces and homogenized on ice. The homogenate was centrifuged at 800g 4 °C for 10 min. The supernatant was transferred to a new centrifuge tube and centrifuged at 10,000g 4 °C for 10 min. The resulting pellets were the crude mitochondrial fraction. The supernatant was centrifuged at 100,000g 4 °C for 60 min. The final supernatant was the pure cytosolic fraction.
The crude mitochondrial fractions were suspended in 1.5 ml 25% nycodenz buffer, layered onto 1.25 ml 30% nycodenz buffer, and overlaid with 1.25 ml 23% nycodenz buffer containing 5 mM Tris, 3 mM KCl, 0.3 mM EDTA, and pH7.5. The samples were centrifuged at 52,000 for 90 min at 4 °C in a swinging bucket rotor (BECKMAN, SW60 Ti). The mitochondrial fraction was collected from the 25%/30% interface and resuspended in equal volume of homogenization buffer and centrifuged at 10,000g at 4 °C for 10 min. This step was repeated for three times. The pure mitochondria fraction in the final pellet was used for the immunoblot assay.
Protein concentrations were determined by BCA protein assay (Thermo Scientific). Equal mass protein samples (10 μg) were subjected to 10% SDS-polyacrylamide gel electrophoresis, transferred to PVDF membrane (MILIPORE), and immunoblotted with primary antibodies. The antibodies used were anti-Cyclophilin F (Abcam, ab110324), 1:1000 dilution; anti-COX-IV (Cell Signaling, 4844S), 1:5000 dilution and anti-GAPDH (Cell signaling, 5174S), 1:10000 dilution. Results were visualized with ECL reagents (Thermo Scientific). Densitometry evaluation was conducted using ImageJ software (NIH, Bethesda, MD).
Statistical comparisons were done using students t test for single mean or ANOVA test for multiple means when appropriate. All graphs were plotted in Sigmaplot (Systat Software Inc.) and results were expressed as mean ± S.E., and p < 0.05 was considered significantly different.
Denervation leads to drastic increase of mitoflash signal in skeletal muscle fibers
Based on the biochemical study of Muller et al. , it has been shown that the ROS production increases 30-fold in skeletal muscle after 7 days of denervation. The initial response of mitochondria to denervation in skeletal muscle has not been studied. The mt-cpYFP transgenic mouse model provides a useful tool to examine the dynamic changes of mitochondrial ROS production in the form of mitoflash signaling in live skeletal muscle cells [25, 26]. While there is ongoing debate regarding the mt-cpYFP sensitivity to ROS and pH , resolution of the spatial and temporal aspects of mitoflash signals has been widely used to explore the changes in the metabolic function of mitochondria under physiological and pathological conditions .
The mt-cpYFP transgenic mice were generated in our laboratory. The mitochondrial targeting of mt-cpYFP is demonstrated in the Additional file 1: Figure S1. The FDB muscle fibers derived from the mt-cpYFP mice were isolated for live cell imaging to record the mitoflash activity. A standard protocol was established to record the mitoflash events in FDB muscle fibers, in which 100 images were taken continuously at a speed of 1 image/s. The mitoflash signal was analyzed using the established software, FlashSniper , to obtain parameters of the mitoflash signal including full area at half maximum (FAHM), full duration at half maximum (FDHM), and the amplitude of the signal (dF/F0). In addition, the fiber area giving mitoflash signal during the 100-sequential-imaging time period was summed as Total Flash Area. The ratio of Total Flash Area over the whole fiber area named Total Flash Area/Fiber Area was then calculated. Total Flash Area/Fiber area during this 100-sequential-image recording time period provides quantification of the number of mitochondria in a single muscle fiber that are involved in generating mitoflash events, as well as the frequency of the mitoflash events.
Additional file 2: movie 1. Mitoflash recording of a denervated muscle fiber; 012Den4231_movie1. (AVI 1458 kb)
Additional file 3: movie 2. Mitoflash recording of a fiber with the sham procedure; 009CL4413_movie2. (AVI 1182 kb)
Cyclosporine A attenuates the increase of mitoflash activity caused by denervation
It has been shown that mitoflash signal is associated with transient opening of the mitochondrial permeability transition pore (mPTP) in both cardiac and skeletal muscle [26, 27]. However, two published studies reported that CsA did not affect the mitoflash signal in normal skeletal muscle [36, 37], suggesting the possibility that the mitoflash signal is not related to Cyclophilin D (CypD) under physiological conditions. Here, we tested whether mitoflash activity became CypD-dependent upon denervation. We examined whether altered mPTP opening was associated with the generation of massive mitoflash signal in denervated skeletal muscle and whether inhibition of CypD-dependent pore opening could restore mitoflash activity to the normal level in denervated muscles. Muscle fibers isolated from denervated skeletal muscle were treated with 1 μM Cyclosporine A (CsA) for 30 min prior to imaging. Additional file 4 (movie 3) and Fig. 1c1–3 show representative images of denervated muscle treated with CsA. The application of CsA significantly decreased the flashing area (Total Flash Area/Fiber Area) in denervated muscle fibers (Fig. 1d; Den 7.77 ± 1.87, n = 59 muscle fibers vs Den + CsA 2.50 ± 0.63, n = 57 muscle fibers; p < 0.001). In line with those results, the size of mitoflash signal (FAHM) was also significantly reduced (Fig. 1e; Den 83.43 ± 19.14, n = 59 muscle fibers vs Den + CsA 50.45 ± 14.96, n = 57 muscle fibers, p = 0.023). Interestingly, the flash amplitude dF/F0 was significantly reduced after the CsA treatment (Fig. 1f; Den 0.96 ± 0.09, n = 59 muscle fibers, Den + CsA 0.79 ± 0.10, n = 57 muscle fibers; p = 0.02). No change in FDHM was observed among the groups. These results indicate that the opening of mPTP plays an important role in the mitochondrial ROS production in the denervated skeletal muscle cells.
Additional file 4: movie 3. Mitoflash recording of a denervated fiber in the presence of CsA; 004CsA72513_movie3. (AVI 976 kb)
Because CypD plays a critical role in controlling mPTP activity, we conducted immunoblot analysis to evaluate the protein expression level of CypD in both mitochondria and cytosol of the skeletal muscle following 24 h of denervation. We first confirmed the purity of mitochondrial and cytosol fractionation by using specific markers for mitochondria (COX-IV) and cytosol (GAPDH). As demonstrated in Additional file 5: Figure S2, the pure mitochondria fractions do not contain GAPDH, while the cytosol portion does not contain COX-IV, indicating no cross-contamination between the mitochondrial and cytosol fractionations. The level of CypD in mitochondria (CypD-mito) and cytosol (CypD-cyto) was quantified by normalization to the mitochondrial and cytosol specific markers, respectively, prior to the calculation of the CypD-mito to CypD-cyto ratio. We found that the ratio of CypD level in mitochondria to the CypD level in cytosol (CypD-mito/CypD-cyto) was significantly increased in the denervated muscle compared with the sham-treated muscle (sham 1.28 ± 0.20 vs Den 2.12 ± 0.31, n = 5, p < 0.05) (Fig. 1g and h). The data provide additional support that the denervation-induced mitoflash signal is likely related to CypD-dependent pore opening of the mPTP.
Electric stimulation reduces mitoflash activity in both sham and denervated muscle fibers
Additional file 6: Movie 4. Mitoflash recording of a denervated muscle fiber before electric stimulation; 005BeforeDenStim050614_movie4. (AVI 1634 kb)
Additional file 7: Movie 5. Mitoflash recording of a denervated muscle fiber 10 s after the electric stimulation; 005aDenStim050614_movie5. (AVI 1334 kb)
Electrical stimulation and CsA treatment reduces the recurrence of mitoflash events in denervated skeletal muscle
Mitochondria Ca2+ transient follows the physiological intracellular Ca2+ transient
The patch pipette solution contained x-rhod-1, a fast cytosolic Ca2+ indicator with excitation and emission spectra distinct from those of mt11-YC3.6. The x-rhod-1 delivered into the cytosol of the muscle fiber (Fig. 5b) allows the recording of Ca2+ transients in the cytosol simultaneously with mitochondrial signal recording of mt-YC3.6 (Fig. 5c, f 3). The change in fluorescence intensity F/F 0 (t) of x-rhod-1 (Fig. 5d, red trace) was used to derive the intracellular Ca2+ transient, and the changes in mito ratio f 3 (f 2/f 1) (Fig. 5d, black trace) was used to derive the mitochondrial Ca2+ uptake using the same method described in our previous work  (Fig. 5e). Figure 5e demonstrates that the dynamic change of Ca2+ level in mitochondria (Fig. 5e, black trace) indeed follows the dynamic change of Ca2+ level in the cytosol (Fig. 5e, red trace). Following a short (20 ms) depolarization stimulation, the time courses of Ca2+ transients are very similar in both mitochondria and cytosol.
Using MitoSOX Red to evaluate the role of intracellular Ca2+ transients on mitochondrial ROS production
Since our voltage-clamp data demonstrated that Ca2+ level in mitochondria followed the dynamic changes of cytosolic Ca2+ level and Ru360 blocked the effect of electric stimulation on mitoflash activity (Fig. 3), we hypothesize that mitochondrial Ca2+ uptake may affect the superoxide production during a physiological Ca2+ release transient. As in Fig. 3, we further tested whether pharmacological inhibition of the mitochondrial Ca2+ uptake by Ru360 could block the effect of the field stimulation on the denervated muscle fibers labeled with MitoSOX Red. The denervated FDB muscle fibers were first incubated with 10 μM Ru360 for 30 min before receiving the field stimulation. Immediately, following the field stimulation (still in the presence of 10 μM Ru360), the denervated FDB fibers were loaded with MitoSOX Red for evaluation of mitochondrial superoxide level. As shown in Fig. 7a4 and b, in the presence of Ru360, the field stimulation no longer reversed the elevated mitochondrial superoxide level in the denervated muscle fibers (15.15 ± 5.16, n = 5, p < 0.05 compared to the denervated fibers without Ru360), suggesting that the transient mitochondrial Ca2+ uptake is likely responsible for suppressing the mitochondrial ROS production during a physiological Ca2+ release transient. We also conducted a control experiment to test if Ru360 has a potential effect on mitochondrial ROS production. As demonstrated in Fig. 7a1 and a2, Ru360 has no significant influence on mitochondrial superoxide production level.
Previously published studies on how Ca2+ signaling regulate mitochondrial ROS production in skeletal muscle mainly focused on the effect of a steady state level of Ca2+ inside mitochondria . Here, we provide the first evidence that the dynamic uptake of Ca2+ by mitochondria may also be a key player in maintaining the physiological function of mitochondria. We showed that the time course of free Ca2+ level inside mitochondria follows the time course of cytosolic Ca2+ transient and that physiological changes in mitochondrial Ca2+ levels could be essential for maintaining the functional integrity of mitochondria. Following denervation, there is no action potential initiated in muscle fibers, and therefore, no Ca2+ transients in the cytosol and mitochondria. Mitochondria respond to this condition with increased ROS production that is associated with mPTP opening. Restoring the physiological Ca2+ transients by electric stimulation reduced the mitochondrial ROS production and stopped the repetitive mPTP opening in the denervated muscle fibers.
Motor neuron innervation is critical for the growth and maintenance of muscle fibers. Denervation is known to cause muscle atrophy, manifested in age-dependent sarcopenia and other neurological diseases, such as amyotrophic lateral sclerosis (ALS). Studies by other investigators have shown that denervation leads to compromised EC coupling machinery in skeletal muscle  and altered mitochondrial metabolic function [42, 43]. The increase in ROS generation is a common event in skeletal muscle mitochondria under a variety of pathological conditions associated with denervation-induced muscle atrophy . Excessive ROS generation contributes to apoptotic or necrotic cell death . Biochemical assay has revealed that the ROS generation in muscle mitochondria was dramatically increased after several days of surgical sciatic nerve transection [10, 44]. It is believed that enhanced ROS generation is a common factor in the mechanism underlying denervation-induced muscle atrophy and the related downstream signaling pathways have been extensively studied [2, 11]. However, the initial signal that causes changes in the mitochondrial network and ROS production of skeletal muscle in response to denervation remains unknown. It is not clear whether the dynamic changes of the Ca2+ level in the cytosol and mitochondria are implicated in this process. Here, we provide evidence that physiological Ca2+ transient is required to maintain the integrity of mitochondrial function in the physiological condition. Absence of the physiological Ca2+ transients is likely a direct cause of enhanced mitochondrial ROS production that is associated with the repetitive opening of the mitochondrial mPTP.
Wang et al. developed a mitochondrial-targeted, circularly-permutated yellow fluorescent protein (mt-cpYFP) as an indicator of ROS production and energy metabolism of individual mitochondrion in live cells including skeletal muscle cells [25–27]. Their studies revealed that ROS is produced in transient waves at the level of individual mitochondria, a phenomenon known as the mitoflash . Using the same transgene, we produced the mt-cpYFP transgenic mice in our laboratory and investigated the effect of denervation on mitochondrial function by monitoring the dynamic changes of mitoflash signals in mitochondria of live muscle fibers. Twenty-four hours following the sciatic nerve transection, denervated muscle fibers show an excessive increase in the mitoflash signal, specifically in the signal size (FAHM) that was four times larger than in control fibers. The data indicate that in a single muscle fiber, more mitochondria are activated to produce mitoflash signals in response to denervation. In addition, the Total Flash Area/Fiber Area of the mitoflash signal recorded during the 100-sequential-imaging time period was four times more than the controls, indicating a higher frequency of individual mitochondria to produce mitoflash activity.
In both skeletal and cardiac muscle cells, mitochondria form a structural and functional network [30, 45, 46]. Imaging of mitoflash signal of skeletal muscle in mt-cpYFP mice has also revealed functional mitochondrial network . In the current study, the dramatic increase in the size of the mitoflash signal (FAHM) could indicate that the mitochondrial network is physically more connected following denervation. However, this is unlikely, as a study by Romanello et al. has reported that denervation promotes mitochondrial fission activity, which should reduce the physical connection between mitochondria . Our previous study has also identified a reduced mitochondrial network with enhanced fission activity in skeletal muscle of an ALS mouse model (G93A), in which skeletal muscle experiences denervation during ALS progression . As identified in cardiac myocytes, the release of ROS from a subset of mitochondria can trigger ROS release from the adjacent mitochondria [48–50]. The dramatically enhanced size of mitoflash signal in skeletal muscle is likely an indication of the ROS-induced ROS release in mitochondria that is augmented by denervation. Other kinetic parameters such as the signal amplitude and FDHM stay the same, indicating that mitoflash kinetics was not affected by a short period (24 h) of denervation.
While mitochondria take up Ca2+ during skeletal muscle contraction [19, 20, 51], the mitochondrial Ca2+ uptake seems to be a double-edged sword for the fate of cells [22, 40]. Under normal conditions, mitochondrial Ca2+ uptake is a physiological stimulus for ATP synthesis [21, 52, 53]. The elevated mitochondrial Ca2+ level leads to a coordinated upregulation of oxidative phosphorylation machinery, resulting in higher mitochondrial ATP output to meet the energy demanding of the cells [54, 55]. This is required to meet the increased contractile force during muscle contraction. In addition, the mouse model (MCU−/−) with global knockout of mitochondrial calcium uniporter (MCU) showed a smaller body size and impaired skeletal muscle performance along with absence of mitochondrial Ca2+ uptake in isolated skeletal muscle mitochondria, indicating that mitochondrial Ca2+ uptake plays an important role in skeletal muscle development and performance . However, Ca2+ overload in mitochondria is also a pathological stimulus of ROS generation . It has been shown that prolonged skeletal muscle inactivity (such as muscle disuse) leads to an increased resting cytosolic free Ca2+ level, that in turn overloads mitochondria to stimulate the ROS production [23, 24]. While previous published studies on how Ca2+ signaling regulate mitochondrial ROS production in skeletal muscle mainly focused on the effect of a steady state level of Ca2+ inside mitochondria, it is not known whether the dynamic Ca2+ uptake by mitochondria has a role in maintaining the mitochondrial functional integrity. Recording of the mitoflash signal in a skeletal muscle fiber following a short period of denervation allowed us to discover the early response of mitochondria to the absence of physiological cytosolic Ca2+ transients, which limits mitochondrial Ca2+ uptake and leads to an excessive increase in mitochondrial ROS production. Our result is in line with the study by Csordas et al. , in which they found that destruction of the ER-mitochondria linkage causes the specific loss of the IP3R-mediated Ca2+ transfer to mitochondria that stimulates oxidative metabolism in a cultured mammalian cell line . Most importantly, we demonstrated that the early increase of mitochondrial ROS production was immediately diminished by restoring a train of physiological intracellular Ca2+ transients in denervated muscle fibers and that the transient mitochondrial Ca2+ uptake could be a key regulator of mitochondrial mPTP activity. Interestingly, recent studies from other research groups also provided evidence of MCU-dependent mitochondrial Ca2+ uptake in protecting denervation-induced skeletal muscle atrophy by using virus-mediated overexpression of MCU [58, 59]. While it is very well known that Ca2+ overload into mitochondria leads to mPTP opening and ROS production, one of the major findings in the present study is that the denervation-induced ROS elevation could be reduced by electrical stimulation, indicating an unexpected role of the Ca2+ uptake into mitochondria for control of ROS production. While a consistently elevated intracellular Ca2+ level leads to the Ca2+ overload in mitochondria and mPTP opening, the absence of physiological Ca2+ transients following denervation, which leads to the absence of the dynamic Ca2+ uptake into mitochondria, also triggers mPTP opening and initiates mitochondrial dysfunction. We speculate that there may be a biphasic dependence of mPTP on Ca2+ level or direct response to SR Ca2+ release. The molecular composition of mPTP is still incompletely understood. It is unclear what are the molecular mechanisms underlying the different responses of mitochondria to a consistently elevated intracellular Ca2+ level and to the absence of physiological Ca2+ transients following denervation. Future studies are needed to further understand the molecular mechanism underlying the initial response of mitochondria to denervation in skeletal muscle.
While denervation eliminates neuromuscular transmission and myoplasmic Ca2+ transients, other muscle adaptations and changes can occur during the first 24 h after denervation. One possibility is that mitochondrial and/or cellular ROS detoxification mechanisms (e.g., SOD, GSH reductase, catalase, and thioredoxin) may be reduced after denervation, and thus, allow for aberrant cell-wide propagation of otherwise spatially restricted mitochondrial ROS-related events. If bursts of ROS-related mitoflash activity are initiated by low-level increases in ROS, then a breakdown of cellular ROS detoxification mechanisms following denervation could also have a contribution to the observed increase in cell-wide waves of ROS-induced ROS release. Thus, changes to denervated fibers in addition to the loss of cytoplasmic Ca2+ transients and mitochondrial Ca2+ uptake are needed to explain the observed enhancement in mitoflash activity. More studies are needed to further understand the molecular mechanism for early response of skeletal muscle to denervation.
One concern about our result is the possibility that the drastic reduction of the mitoflash signal in the denervated muscle fibers following the tetanic field stimulation is due to the cell damage caused by the tetanic field stimulation. We were aware of the study of mitoflash signal on normal skeletal muscle by Wei and colleagues who reported that the mitoflash frequency was significantly increased following a 2-s tetanic stimulation and markedly decreased following prolonged 20-s tetanic stimulation . As a result, the rationale for our experiment design is to avoid the 2-sec tetanic stimulation that already showed promoting mitoflash activity. We also decided not to use 20-s stimulation, as we do not know if the 20-s tetanic stimulation could cause artificial responses in our study. Instead, we applied a tetanic stimulation with much shorter duration (350 ms). Thus, the reduction in mitoflash signal observed in the denervated muscle fibers following such a brief electrical stimulation is unlikely due to the physical damage caused by the field stimulation. Interestingly, this brief stimulation also suppressed the mitoflash activity in the sham muscle. Our data together with the previous study by Wei et al., suggest that the response of mitochondria to tetanic stimulation is dynamic, and the underlying molecular mechanisms may be different between the 350-ms stimulation and 2-s stimulation. It is not a surprise to see that a sevenfold longer electric stimulation (2 s) enhances the mitoflash activity, as it likely promotes more mitochondrial Ca2+ uptake, which is known to promote the function of the respiratory chain reaction.
We previously demonstrated that mitochondria in skeletal muscle take up Ca2+ following a brief electric stimulation . Here, we showed that during prolonged membrane depolarization, mitochondrial Ca2+ transient also follows a similar time course as the cytosolic Ca2+ transient in skeletal muscle fibers. Our results support the hypothesis that the dynamic levels of Ca2+ inside mitochondria follow the time course of the cytosolic Ca2+ transients initiated by motor neuron activation in the physiological condition and that the dynamic change in mitochondrial Ca2+ level could serve as a sensor to decode the nerve innervation. While Ca2+ overload in mitochondria triggers more ROS production , our results indicate that absence of the physiological mitochondrial Ca2+ uptake also play a role for maintaining the integrity of normal mitochondrial function.
The coupling of mitoflash and mPTP opening has been well characterized in both cardiac and skeletal muscle [27, 35]. It has been shown that mt-cpYFP fluorescence is transiently increased when the mitochondrial membrane potential (measured by TMRE) is depolarized during a dual recording, indicating a dynamic coupling of the mitoflash event and the mPTP opening. Several lines of evidence have suggested a strong connection between CypD-related mPTP opening and the mitoflash signal in cardiac muscle but not in normal skeletal muscle. Indeed, manipulation of mPTP had a major impact on the mitoflash signal in cardiac myocytes. For example, activators of mPTP, like atractyloside, increased the mitoflash frequency in cardiac myocytes  but not in skeletal muscle fibers . In contrast, CypD inhibitors such as cyclosporine A (CsA) caused a reduction in the mitoflash amplitude and kinetics in cardiac myocytes . However, CsA did not appear to affect the mitoflash signal in skeletal muscle fibers under normal physiological conditions [36, 37], indicating that the mitoflash signal in normal skeletal muscle may not be related to the activity of CypD at physiological conditions. Remarkably, we demonstrated that in the case of denervation, the enhanced mitoflash signal might directly associated with the CypD-related opening of mPTP in skeletal muscle fibers. Incubation with 1 μM CsA for 30 min significantly reduced the Total Flash Area/Fiber Area, FAHM, and the amplitude of the mitoflash in denervated muscle fibers. CsA inhibits mPTP opening by binding to Cyclophilin D (CypD) . Thus, the apparent relationship between mPTP and the mitoflash signal seems to be related on CypD in the denervated muscle. It has been shown that knocking down CypD with siRNA, a major component related to mPTP, significantly reduced mitoflash frequency in cardiac myocytes [26, 61]. The opposite occurred upon CypD overexpression . Based on a biochemical study, Csukly K et al. has shown that following 3 weeks of denervation, the isolated mitochondria from denervated muscle have enhanced vulnerability to Ca2+-induced mPTP opening, and this Ca2+-induced mPTP opening can be blocked by the application of CsA. In addition, the relative level of CypD is increased in mitochondrial fraction of the denervated skeletal muscle, indicating a CypD-related mPTP opening in denervated muscle . In line with this discovery, we found that 24-h denervation led to an increase in the ratio of CypD-mito/CypD-cyto. The increased ratio of CypD-mito/CypD-cyto could reflect increased degradation of the cytosolic CypD, increased stability of mitochondrial CypD, or increased translocation of CypD from the cytosol to mitochondria. Nevertheless, the increased ratio indicates a relative change of CypD in mitochondria fraction. This result together with the pharmacological study of CsA suggests that CypD is likely involved in regulating mPTP activity and mitochondrial ROS production in skeletal muscle following 1 day of denervation. Although it is very well established that CsA inhibits mPTP by binding to CypD and has been extensively used as an investigative tool for mPTP research, it is also known that CsA inhibits calcineurin, a calcium dependent dephosphorylation enzyme [60, 62]. Studies by Cereghetti et al. and Cribbs and Strack demonstrate the role of calcineurin in dephosphorylation of Drp1, which promotes mitochondrial fission [63, 64]. It has been found that CsA promotes mitochondrial fusion through inhibition of calcineurin [63, 65]. Thus, it is possible that the reduced mitoflash activity in the presence of CsA may also partially due to the enhanced mitochondrial fusion, which indirectly affects the mitochondrial ROS production. Nevertheless, our result with relatively enhanced mitochondrial CypD level and the published study showing an enhanced CypD expression level in skeletal muscle with prolonged denervation suggest a potential involvement of the CypD-related mPTP opening in skeletal muscle in response to denervation. The CypD-related mPTP opening is likely an early event in skeletal muscle mitochondria in response to denervation. Most importantly, for the first time, our data demonstrated that the physiological Ca2+ transients and the following mitochondrial Ca2+ uptake play a key role in maintaining the functional integrity of mitochondria through a potential mechanism by regulating the mPTP-related mitochondrial ROS production.
Although mt-cpYFP has been characterized and used as a biomarker of mitochondrial ROS generation [26, 36, 37], there are concerns about the specificity of the mitoflash signal. It has been suggested that mitoflash may also report ATP  or pH changes inside mitochondrial matrix [34, 67]. Nevertheless, the mitoflash signal may reflect the status of the ROS-related metabolic function of mitochondria, and cpYFP likely can report both ROS and pH signal of mitochondria, and is thus a robust biosensor for mitochondrial function [68, 69]. However, the portion of ROS vs pH signal reported by the mitoflash signal could be context-dependent . Such detailed quantification will require additional measurement, which is not the focus of the current study. A recent study by Ding et al. indicates that mitoflash represents a dynamic signal of mitochondrial ROS production and energy metabolism . In our current study, mitoflash signal served as a unique biosensor that allowed us to dissect the early response of mitochondria to denervation and to identify the role of physiological Ca2+ transients in maintaining the functional integrity of mitochondria in skeletal muscle. Due to the ongoing debate of mt-cpYFP sensitivity on ROS and pH , we used MitoSOX Red, a separate fluorescent indicator for mitochondrial superoxide detection, to further evaluate the response of denervated muscle fibers to electrical stimulation. Although MitoSOX Red is not able to detect the dynamic changes of mitochondrial ROS production, its fluorescence intensity can report the basal level of mitochondrial superoxide production. Our MitoSOX Red data confirmed that mitochondrial ROS production was enhanced following 24 h of denervation and that electrical stimulation could reverse mitochondrial ROS production to basal level.
In summary, we have taken advantage of the transgenic mt-cpYFP mice that allowed us to evaluate the early dynamic response of skeletal muscle to denervation at the mitochondrial level of live muscle fibers. For the first time, our study reveals that physiological Ca2+ transient is important for maintaining the normal mitochondrial metabolic function by regulating the mPTP-associated mitochondrial ROS generation. Combining voltage-clamp technique and live skeletal muscle cell imaging study, we showed that the Ca2+ level inside mitochondria of skeletal muscle is synchronized with the dynamic change of cytosolic Ca2+ level under physiological condition. In the case of denervation, there are no Ca2+ transients initiated in the cytosol and thus no dynamic alterations in mitochondrial Ca2+ level. Our study suggests that the dynamic change of Ca2+ level inside mitochondria regulates the mitochondrial ROS generation as a response to the cytosolic Ca2+ transient in the physiological condition. Additionally, mitochondria in denervated muscle fibers could sense the absence of the physiological Ca2+ transients and respond with enhanced ROS generation that could initiate downstream signaling toward mitochondrial dysfunction. Future studies should focus on the detailed molecular basis underlying the mitochondrial response to the absence of physiological Ca2+ transients.
Amyotrophic lateral sclerosis
- E-C coupling:
Full amplitude of half maximum
Flexor digitorum brevis
Full duration of half maximum
Mitochondrial transition pore
Mitochondrial targeted biosensor
Reactive oxygen species
We appreciate Mr. Frank Yi for editing this manuscript.
This work was fully supported by Muscular Dystrophy Association Grant MDA-4351 and NIAMS/National Institutes of Health Grant R01 AR057404 to JZ and partially supported by R01-AG028614 to JM. CK was a recipient of a postdoctoral fellowship from NIH/NHLBI T32 training grant HL 07692-(21-25). LZ was a recipient of a scholarship from Zunyi Medical University. JZ laboratory is also supported by Victor E. Speas Foundation and McCown Gordon Gala Research Gift.
Availability of data and materials
All data generated or analyzed during this study are included in this published article.
JZ designed and supervised the study. JM and JY participated the experimental design of the study. JZ, JM, CK, and JY wrote the manuscript. CK, JY, YX, KD, JZ, XL, and CM performed the experiments. CK, KD, YX, JY, JM, and JZ analyzed the data. KL and JX assisted running the image-process software. HC participated in the scientific discussion on this study. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
Consent for publication
All animal experiments were performed according to the procedures approved by Institutional Animal Care and Use Committee of Rush University Medical Center, University of Missouri at Kansas City, and Kansas City University.
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