Substrate-dependent and cyclophilin D-independent regulation of mitochondrial flashes in skeletal and cardiac muscle

Abstract

Mitochondrial flashes (mitoflashes) are stochastic events in the mitochondrial matrix detected by mitochondrial-targeted cpYFP (mt-cpYFP). Mitoflashes are quantal bursts of reactive oxygen species (ROS) production accompanied by modest matrix alkalinization and depolarization of the mitochondrial membrane potential. Mitoflashes are fundamental events present in a wide range of cell types. To date, the precise mechanisms for mitoflash generation and termination remain elusive. Transient opening of the mitochondrial membrane permeability transition pore (mPTP) during a mitoflash is proposed to account for the mitochondrial membrane potential depolarization. Here, we set out to compare the tissue-specific effects of cyclophilin D (CypD)-deficiency and mitochondrial substrates on mitoflash activity in skeletal and cardiac muscle. In contrast to previous reports, we found that CypD knockout did not alter the mitoflash frequency or other mitoflash properties in acutely isolated cardiac myocytes, skeletal muscle fibers, or isolated mitochondria from skeletal muscle and the heart. However, in skeletal muscle fibers, CypD deficiency resulted in a parallel increase in both activity-dependent mitochondrial Ca²⁺ uptake and activity-dependent mitoflash activity. Increases in both mitochondrial Ca²⁺ uptake and mitoflash activity following electrical stimulation were abolished by inhibition of mitochondrial Ca²⁺ uptake. We also found that mitoflash frequency and amplitude differ greatly between intact skeletal muscle fibers and cardiac myocytes, but that this difference is absent in isolated mitochondria. We propose that this difference may be due, in part, to differences in substrate availability in intact skeletal muscle fibers (primarily glycolytic) and cardiac myocytes (largely oxidative). Overall, we find that CypD does not contribute significantly in mitoflash biogenesis under basal conditions in skeletal and cardiac muscle, but does regulate mitoflash events during muscle activity. In addition, tissue-dependent differences in mitoflash frequency are strongly regulated by mitochondrial substrate availability.

INTRODUCTION

Mitochondrial flashes (mitoflashes) are stochastic events of changes in mitochondrial superoxide, pH, and, membrane potential (Ψ_m) detected with mitochondrial-targeted cpYFP (mtcpYFP) across a wide range of species and cell types [1–6]. Mitoflashes were initially proposed to reflect quantal events of mitochondrial superoxide production given the unique sensitivity of purified cpYFP to superoxide and not to other forms of reactive oxygen species (ROS), ATP, NAD+/NADH, or redox potential. However, due to the intrinsic pH sensitivity of GFP-derived probes, whether mitoflashes represent changes in superoxide, matrix alkalinization, or both, is hotly debated [7–10]. Simultaneous measurements of mitoflashes (using mt-cpYFP) and matrix pH (using SNARF-1) revealed that the mitoflash events primarily reflect a burst in superoxide that coincides with a modest degree of Ψ_m depolarization and matrix alkalinization [11, 12].

Subsequent studies proposed that the coincidence of mitochondrial ROS production, pH alkalinization, and Ψ_m depolarization are intimately linked to mitochondrial metabolism [13, 14]. Mitoflash activity is tightly coupled to mitochondrial respiration, requires functional electron transport chain (ETC) activity, and a robust proton motive force across the mitochondrial inner membrane [1, 11]. Physiologically, mitoflash events are a highly-sensitive biomarker of mitochondrial metabolism regulated by cellular levels of ROS, Ca²⁺ and protons [3, 6, 15–18]. Pathologically, mitoflash activity is increased in several oxidative-stress related disorders, including ischemia reperfusion [1], malignant hyperthermia [3, 19], ROS-induced apoptosis [20] and muscular dystrophy [6]. However, the precise mechanism(s) that control mitoflash generation and termination remain to be fully elucidated [21].

The depolarization of Ψ_m during a mitoflash event results from the opening of a non-selective, large pore channel within the mitochondria inner membrane [1]. The mitochondrial membrane permeability transition pore (mPTP) is a large conductance channel that, once activated, allows molecules smaller than ~1.5 kDa to pass freely across the mitochondrial inner membrane. Although the molecular identity and specific mechanism for its activation have yet to be fully defined, the mPTP is considered to be the most likely candidate for the large conductance pore activated during a mitoflash. Supporting this idea, the mPTP activator atractyloside increases mitoflash frequency, while knocking out the mPTP regulator cyclophilin D (CypD) reduces mitoflash frequency in cardiac myocytes [1, 22]. However, this effect of CypD on mitoflash activity appears to be tissue-dependent as CypD ablation in skeletal muscle does not significantly alter mitoflash activity either in-vivo or in-vitro [3, 22]. Interestingly, the expression of CypD protein in the heart is 4-fold higher than that observed in skeletal muscle [22].

In this study, we set out to further investigate the tissue-specific effects of CypD and mitochondrial substrates on mitoflash activity in skeletal and cardiac muscle. We took advantage of the fact that one of our mt-cpYFP transgenic mouse lines exhibits significant mtcpYFP expression in both skeletal and cardiac muscle. Unexpectedly, we were unable to resolve a tissue-specific effect of CypD ablation on mitoflash activity between skeletal and cardiac muscle in either intact cells or isolated mitochondria. Instead, we found that CypD was essential for activity-dependent regulation of mitoflash activity in skeletal muscle. We also compared the tissue-specific effects of different mitochondrial substrates on mitoflash activity in both intact cells and isolated mitochondria from skeletal and cardiac muscle and identified a tissue-specific regulation of mitoflash activity by mitochondrial substrates in intact skeletal and cardiac muscle cells.

METHODS

Animals

Mt-cpYFP/CypD KO compound mice were generated by crossing muscle-specific, mt-cpYFP transgenic mice [3] with CypD KO mice. F2 generation mice were used for experiments with CypD KO and WT littermates expressing mt-cpYFP. Mice were housed at the animal facility at the University of Rochester School of Medicine and Dentistry, Rochester, NY. All animal protocols were approved by the University of Rochester Committee on Animal Resources. Mice were group-housed in sterile ventilated micro-isolator cages on corn cob bedding in an AAALAC accredited facility. Animals were provided ad libitum access to pelleted feed (LabDiet 5010) and water (Standard drinking water of Rochester, NY, pH 7.8) via HYDROPAC®. Animals were maintained on a 12:12 hour light:dark cycle in rooms at 72°F with 30–70% humidity under pathogen-free conditions. Age-matched littermates were used for all experiments and all data sets included male and female mice.

When the muscle–specific, mt-cpYFP transgenic mice were created, several lines of mice were established from different founders. Line 96 expresses mt-cpYFP specifically in skeletal muscle. However, line 18 was found to express mt-cpYFP in both skeletal muscles and the heart, but not in other tissues. The majority of the experiments conducted in this study used line 96 for skeletal muscle and line 18 for cardiac myocytes. However, for comparison of mitoflash properties between skeletal and cardiac muscles, only line 18 mice were used for both tissue types.

Isolation of mitochondria from skeletal muscle

Mitochondria from skeletal muscle were isolated as described previously [11]. Briefly, hindlimb muscle tissue from control mt-cpYFP and mt-cpYFP/CypD KO mice were dissected on ice in Chappell-Perry (CP) buffer containing 50 mM Tris, pH 7.4, 100 mM KCl, 5 mM MgCl₂ and 1 mM EDTA, and manually minced using scissors. Minced muscle tissue was washed twice in CP buffer, digested on ice for 6 min in 1 mg/ml proteases (from Streptomyces griseus Type XIV, Sigma) and homogenized in a potter homogenizer equipped with a motor driven pestle (4 strokes at 500 rpm). The supernatant was collected following a low speed centrifugation at 3500 × g for 4 min at 4°C. Digestion and centrifugation were repeated once with the pellet and supernatant collected as above. The pellet was exposed to a final 8 min digestion at room temperature followed by centrifugation. The supernatants from each centrifugation were pooled and subjected to a high-speed centrifugation at 11,000 × g for 9 min at 4°C. The resultant pellet contained the mitochondrial fraction. The pellet was washed once in a buffer containing 50 mM Tris, pH 7.4, 100 mM KCl and 1 mM EDTA. The final pellet was resuspended in skeletal muscle respiration buffer (SKRB, 10 mM HEPES, pH 7.4, 125 mM KCl, 5 mM MgCl₂, 2 mM K₂HPO₄ and 1 mg/ml BSA) without substrates. Various mitochondrial substrates and 40 nM tetramethylrhodamine ethyl ester (TMRE) were added prior to plating on Cell-tak coated coverslips for 30 mins prior to imaging. Mitochondria were stored on ice and used for experiments within 3 hours of preparation.

Isolation of mitochondria from mouse heart

Control mt-cpYFP and mt-cpYFP/CypD KO mice were anesthetized by i.p. injection of 100 mg/kg ketamine, 10 mg/kg xylazine and 2 mg/kg acepromazine. After anesthesia, the heart was removed from the chest and retrogradely perfused through the aorta to remove residue blood using isolation medium (HMIM), containing 300mM sucrose, 2mM EGTA and 20mM Tris, pH 7.35 at 4°C. The heart was then immersed in 2 ml HMIM and homogenized with a tissue homogenizer for 20 strokes (1s per stroke but with the final stroke being 5s). The homogenate was then centrifuged at 1000 × g for 5 mins at 4°C. The resultant supernatant was subjected to a second centrifugation at 10,000 × g for 5 mins at 4°C. The pellet (mitochondrial fraction) was resuspended in cardiac respiration buffer (CRB) containing 120 mM KCl, 5 mM Mg2Cl, 5 mM KH₂PO₄ 100 μM EGTA and 10 mM HEPES, pH7.3. The mitochondrial suspension was stored on ice and used within 1h of isolation. Isolated mitochondria were plated on Cell-tak treated coverslips 30 min prior to imaging in the presence of defined substrates and 40 nM TMRE.

Isolation of cardiac ventricular myocytes

Cardiac ventricular myocytes were isolated from 2–3 months old mt-cpYFP and mt-cpYFP/CypD KO mice. Hearts were quickly excised from anesthetized mice (using anesthetics described in the previous section), cannulated through the aorta, and mounted on a Langendorff perfusion system warmed to 37°C. The heart was first perfused with isolation buffer (in mM: 120 NaCl, 15 KCl, 0.6 Na₂HPO₄, 0.6 KH₂PO₄, 1.2 MgSO₄, 4.6 NaHCO₃, 30 taurine, 5.5 glucose, 10 butanedione monoxime, and 10 HEPES pH 7.4 at 37°C) to remove residual blood from the coronary vessels and then digested at 37°C for 10 mins in isolation buffer supplemented with 12.5 μM CaCl₂, 0.025% trypsin, 0.17U/ml collagenase A, and 0.42U/ml collagenase D. After digestion, the atria and aorta were removed from the heart and the ventricular tissue was crudely separated using fine tweezers. Myocytes were dissociated via trituration using a plastic transfer pipette in stop buffer (isolation buffer with 10% fetal bovine serum (FBS) and 12.5 μM CaCl₂), then filtered through a 200 μm mesh filter and allowed to equilibrate in stop buffer for 10 mins. The resulting pellet was washed once in stop buffer, after which Ca²⁺ was added back to the solution gradually to a final concentration of 1 mM. The myocytes were sedimented by gravity for a final 10 mins. The supernatant was removed and the myocyte pellet was resuspended in culture media MEM supplemented with 10 % FBS, 1 % penicillin and streptomycin (P/S) and 10 μM blebbistatin to inhibit spontaneous contraction. Myocytes were plated on ECL treated coverslips and stored in an incubator at 37°C with 5% CO₂. Myocytes were allowed to equilibrate for 1h before experiments. Prior to imaging, myocytes were transferred to Ringer’s solution with 10 μM blebbistatin and 10 mM glucose or other substrates (Na Pyruvate, Na Palmitate and L-carnitine) where indicated, and with 20 nM TMRE to enable simultaneous monitoring changes in Ψ_m during mitoflash events. Sodium palmitate was conjugated to BSA as per manufacturer instruction (Seahorse bioscience Inc. [23]) in order to increase water solubility. L-carnitine was used with Na-palmitate as a cell membrane transporter.

Isolation of flexor digitorum brevis (FDB) muscle fibers

Single FDB muscle myofibers were isolated using an enzymatic digestion protocol as described previously [3]. FDB muscles from 4-month-old WT and CypD KO mt-cpYFP transgenic mice were digested in 1.2 mg/mL collagenase A in Ringer’s solution for 1 h at 37°C with gentle agitation. Tissue was dissociated by trituration, plated on glass bottom dishes and allowed to settle for 20 min. FDB fibers then were transferred to culture medium (1:1 DMEM/F12, 2 % FBS and 1 % P/S) and kept at 37°C with 5 % CO2 until use. Before experiments, isolated FDB fibers were transferred to Ringer’s solution containing 10 mM glucose (or other substrates where indicated) and 20 nM TMRE for 10 min prior to imaging in order to permit simultaneous monitoring of both mitoflash events and changes in mitochondrial membrane potential.

Confocal imaging and analysis of mitoflashes.

Mitoflash activity was measured as previously described [3, 11]. Briefly, FDB fibers, ventricular cardiac myocytes, and isolated mitochondria from skeletal or cardiac muscle were mounted on the stage of a Nikon Eclipse C1 Plus Confocal microscope equipped with SuperFluor 40× (1.3 NA) oil immersion objective. Mt-cpYFP and TMRE were excited using 488 nm and 543nm lasers and emission detected at 515/30nm and 605/75nm, respectively. Time-lapse x, y image acquisition was obtained using 512×512 resolution, with a total 100 frames, 1.24s/frame (total 124s) for intact cells or 50 frames (total 62s) for isolated mitochondria. Automated detection and analysis of individual mitoflash events during time-lapse x, y imaging were performed using a custom-developed program (“Flash Collector” [3]). Mitoflash frequency, amplitude, full duration at half maximum (FDHM), time constant of decay (τ decay) and spatial area were quantified from x, y time series images.

High-frequency tetanic stimulation of single FDB fibers

FDB fibers isolated from WT and mt-cpYFP/CypD KO mice were bathed in Ringer’s solution supplemented with 25 μM N-benzyl-p-toluene sulfonamide (BTS), a skeletal muscle myosin inhibitor, to inhibit movement during electrical stimulation. Fibers were electrically stimulated with 5 successive tetani (500 ms duration, 100 Hz, 0.2 duty cycle) using an extracellular electrode filled with 200 mM NaCl placed adjacent to the cell of interest [3]. Mitoflash activity was monitored using consecutive x, y time series confocal images before electrical stimulation and immediately after termination of the final tetanic stimulus train. In some experiments, mitochondrial Ca²⁺ uptake was inhibited by incubating fibers with 20 μM Ru360 for 1h before experiments. Some fibers were pre-treated with 1μM cyclosporine A (CsA) 20 minutes prior to experiments to pharmacologically inhibit CypD-dependent mPTP.

Activity-dependent mitochondrial Ca²⁺ uptake measurements

Mitochondrial Ca²⁺ uptake were measured using rhod-2 in FDB fibers following a series of tetanic stimulation as previously described [24]. Briefly, isolated FDB fibers were loaded with 5 μM rhod-2 in Ringer’s solution for 30 min at RT. Minimizing dye and loading time at room temperature facilitated preferential mitochondrial rhod-2 loading. Rhod-2-loaded fibers were bathed in Ringer’s solution supplemented with 25 μM BTS and stimulated using the same paradigm used for activity-dependent mitoflash measurements. Rhod-2 fluorescence was measured before stimulation and right after the 1^st and 5^th tetani. Mitochondrial Ca²⁺ uptake amplitude was calculated as the relative change of rhod-2 fluorescence (F/F₀) in the triad region (I band) after background subtraction using the fluorescence in the non-triad region (A band).

SDS-PAGE and western-blot

Tibialis anterior (TA) muscle and isolated cardiac mitochondria from WT and CypD KO mtcpYFP transgenic mice were used to confirm the ablation of CypD protein in skeletal and cardiac muscle, respectively. TA muscle and cardiac mitochondria were lysed in RIPA buffer, electrophoresed, and transferred to nitrocellulose membrane as described previously [11]. The membrane was incubated with anti-CypD (MitoSciences 1:2000 dilution), anti-MCU (mitochondrial Ca²⁺ uniporter, Abcam 1:2000), anti-mitofusion 1 (Abcam, 1:500 dilution, as loading control) and anti-mitofusion 2 (Sigma, 1:1000, as loading control) antibodies at 4°C overnight, followed by IRDye 800 anti-rabbit secondary antibody (1:10000 dilution) and visualization using a LI-COR imager.

Statistical analyses

Output data from the Flash Collector program were tabulated, averaged, and evaluated for statistical significance using Microsoft Excel and SigmaPlot software suites. Data were expressed as Mean ± S.E.M. Unpaired two-tail Student-t test was used for single comparisons unless stated otherwise. One-way ANOVA followed by Tukey post-hoc test was used for multiple comparisons. A Pearson correlation analysis was used for correlation studies between mitoflash amplitude and duration. P<0.05 was considered statistically significant.

RESULTS

CypD ablation does not alter mitoflash frequency in intact skeletal and cardiac muscle cells

One of our skeletal muscle-specific mt-cpYFP transgenic mouse lines (line 18) was found to express mt-cpYFP in both skeletal muscle and heart (Fig. 1A & B left). We took advantage of this mouse line to study mitoflash properties in the heart and its regulation by CypD by crossing these mice with CypD KO mice. While CypD was previously reported to regulate mitoflash activity in the heart (but not skeletal muscle) [1–3, 22], we found that mitoflash frequency was not significantly different between cardiac ventricular myocytes isolated from WT and CypD KO mice (1.32±0.31 flashes/1000 μm²/100 s versus 1.85±0.38 flashes/1000 μm²/100 s, respectively; Fig. 1A & B right, Fig. 1D). Mitoflash events from WT and CypD KO myocytes also exhibited similar kinetics and occurred concomitantly with mitochondrial membrane depolarization (Fig. 1C), consistent with previous observations [22]. Mitoflash amplitude, FDHM and tau of decay values were also similar in WT and CypD KO cardiac myocytes (Supplemental Fig. 1C & D). Mitoflash frequency (Fig. 1E), magnitude or temporal properties (Supplemental Fig. 1A & B) were also unaltered by CypD deficiency in acutely isolated FDB muscle fibers from another line of mice (line 96) with skeletal muscle specific mt-cpYFP expression, consistent with that reported previously using global mt-cpYFP transgenic mice [22] or following transient expression of mt-cpYFP in FDB muscles of CypD KO mice [3]. Western blot analyses confirmed the absence of CypD protein in both skeletal and cardiac muscle of the CypD KO mice used in these studies (Supplemental Fig. 1E).

Figure 1. Mitoflash activity is unaltered in cardiac myocytes and FDB fibers from CypD KO mice.

(A) Left: representative mt-cpYFP image of a cardiac myocyte isolated from a control mt-cpYFP mouse. Boxed region indicate a region of interest with mitoflash activity. Right: automated standard deviation map generated by “flash collector”. Mitoflash area is circled in magenta. (B) Same as A, except from a CypD KO mouse. (C) Representative traces of mtcpYFP and TMRE fluorescence over time during a mitoflash event within the boxed regions shown in A (left) and B (right). (D) Average (±SEM) mitoflash frequency in cardiac myocytes from WT and CypD KO mice. Inset: same data replotted on an expanded scale. (E) Average (±SEM) mitoflash frequency in FDB muscle fibers from WT and CypD KO mice. n=12–19 cells from 3 mice for cardiac myocytes; n=84–107 cells from 3–4 mice for FDB fibers. Student-t test p>0.05. Scale bars: 10μm.

CypD ablation does not alter mitoflash frequency in mitochondria isolated from skeletal muscle and the heart

Next, we investigated mitoflash activity in isolated skeletal and cardiac mitochondria. Isolated mitochondria were bathed in either SKRB (skeletal) or CRB (cardiac) in the presence of 10 mM succinate as substrate (Complex II). Consistent with results from intact cells, mitoflash frequency in mitochondria isolated from skeletal muscle (Fig. 2A & B) and heart did not differ between WT and CypD KO mice (Fig. 2C & D; Supplemental movies 1 & 2). In addition, mitoflash amplitude and duration were also similar for mitochondria isolated from skeletal or heart of WT and CypD KO mice (Supplemental Fig. 2). Carboxy-atractyloside (CATR), which increases mPTP activity, was previously shown to enhance mitoflash activity in cardiac myocytes from both WT and CypD KO mice [1, 22]. Consistent with this, we found that 10 μM CATR significantly increased mitoflash frequency in isolated cardiac mitochondria from CypD KO mice (Fig. 2E & F). However, CATR did not affect mitoflash amplitude and modestly increased mitoflash duration in isolated cardiac mitochondria from CypD KO mice (Supplemental Fig. 3A & B). We also confirmed that CATR increased mitoflash frequency in intact FDB fibers and cardiac myocytes from WT mice (Supplemental Fig. 3C & D).

Figure 2. Mitoflash activity is unaltered in mitochondria isolated from CypD KO mice.

(A) Surface plot of standard deviation map for mitoflash activity in skeletal muscle mitochondria isolated from either WT (left) or CypD KO (right) mice in the presence of 10 mM succinate. (B) Average (±SEM) mitoflash frequency in skeletal mitochondria isolated from WT and CypD KO mice. Student-t test, p>0.05. n=27 measurements from 4 mice for WT, n=14 measurements from 3 mice for CypD KO. (C-D) Same as A-B, except that mitochondria were isolated from hearts of WT and CypD KO mice. Student-t test, p>0.05. n=39 measurements from 5 mice for WT, n=14 measurements from 3 mice for CypD KO. (E-F) Average (±SEM) mitoflash frequency in mitochondria isolated from hearts of CypD KO mice in the absence (left) and presence (right) of 10 μM carboxyatractyloside (CATR). *p<0.05 compared with control, Student-t test; n=22 measurements from 3 mice for both control and CATR. Mitoflash frequency of the particular measurement is indicated in the bottom right corner of the surface plot.

CypD limits activity-dependent increases in mitoflash activity in FDB fibers

We previously reported that mitoflash frequency in FDB fibers is enhanced after several successive tetanic stimulations [3], presumably due to increased Ca²⁺ uptake into mitochondria during repetitive stimulation. Since CypD-dependent mPTP activity was not required for mitoflash activity under basal conditions, we hypothesized that CypD may regulate mitoflash production during muscle activity through effects on activity-dependent mitochondrial Ca²⁺ accumulation. Indeed, following 5 successive 500ms tetani (at 100Hz), FDB fibers from both control and CypD KO mice exhibited a significant increase in mitoflash activity (Fig. 3A–C), though the relative increase was significantly greater in fibers from CypD KO mice (Fig. 3D). This greater increase in mitoflash activity in CypD KO fibers following electrical stimulation was also observed in control FDB fibers pre-incubated with 1μM CsA, a CypD-dependent mPTP inhibitor, before being subjected to the same stimulation protocol (Fig. 3D). Pre-treatment with 20 mM Ru360, an inhibitor of the mitochondrial Ca²⁺ uniporter (MCU), abolished the activity-dependent increase in mitoflash activity observed in FDB fibers from both WT and CypD KO mice (Fig. 3D), suggesting that the increase in mitoflash activity induced by tetanic electrical stimulation is Ca²⁺ dependent. Average mitoflash amplitude and duration were similar under all conditions (data not shown).

Figure 3. Activity-dependent increase in mitoflash frequency in FDB fibers is further enhanced in CypD KO mice.

(A-B) Representative standard deviation maps of mitoflash frequency before (left) and after (right) 5 consecutive tetanic electrical stimulations in WT (A) and CypD KO (B) FDB fibers. (C) Schematic registry of individual mitoflash events during the 120 s measurements for each condition in A and B. The frequencies for each particular measurement are indicated on the right. (D) Average (±SEM) relative increase in mitoflash frequency after 5 tetanic electrical stimulations (normalized to the frequency before stimulation) for FDB fibers from control and CypD KO mice, after pre-incubation with 1 μM CsA for 20 minutes, and after pre-treatment for 1 hour with 20 mM Ru360. * p<0.05 compared with each corresponding condition before stimulation (red dash line), paired Student-t test. # p<0.05 compared with control after 5 tetani (black bar), one-way ANOVA with Tukey post-hoc test. n=20 cells from 4 mice for control; n=21 cells from 6 mice for CypD KO; n=13 cells from 2 mice for control with CsA; n=7 cells from 2 mice fro CypD KO with CsA; n=8 from 2 mice for control with Ru360; n=9 from 2 mice for CypD KO with Ru360. (E) Representative images of rhod-2 fluorescence for mitochondrial Ca²⁺ uptake before stimulation and after the 5^th tetanus. (F) Average (±SEM) peak amplitude of mitochondrial Ca²⁺ uptake after 5 tetanic electrical stimulations. * p<0.05 compared with WT control; one-way ANOVA with Tukey post-hoc test; n=12–28 cells from 3–5 mice. Scale bars: 10 μm.

To further support our hypothesis that CypD regulates activity–dependent increases in mitoflash activity through mitochondrial Ca²⁺, we measured activity-dependent mitochondrial Ca²⁺ uptake in CypD KO fibers using a Ca²⁺ indicator, rhod-2. A significant increase in mitochondrial Ca²⁺ uptake was detected in FDB fibers from CypD KO mice following electrical stimulation compared to that observed for fibers from WT mice (Fig. 3E & F), in accordance with the greater increase in mitoflash activity observed in fibers from CypD KO mice. Activity-dependent mitochondrial Ca²⁺ uptake was markedly reduced by Ru360 (Fig. 3E & F), consistent with the uptake being mediated by the MCU. Together, these results indicate that CypD regulates mitoflash activity by limiting mitochondrial Ca²⁺ accumulation induced by repetitive, high-frequency stimulation. As a result, an increase in mitochondria Ca²⁺ accumulation following CypD ablation enhances mitochondrial respiration and mitoflash activity. Finally, the increase in activity-dependent mitochondrial Ca²⁺ accumulation in fibers from CypD KO mice is not due to an increase in MCU protein expression (Supplemental Fig. 4), suggesting that CypD promotes mitochondrial Ca²⁺ efflux through activation of the mPTP.

Mitoflash frequency is regulated by cellular environment

An interesting finding from the results presented in Figs. 1 and 2 is that while mitoflash frequency is significantly higher in intact FDB fibers compared to cardiac myocytes (Fig. 1D and E), this difference is lost following mitochondrial isolation in a succinate-containing buffer (Fig. 2B and D). Since the studies in Figs. 1 and 2 were conducted using different mt-cpYFP transgenic lines, we examined this observation more rigorously using a single transgenic line (line 18) in which mt-cpYFP is expressed in both the heart and skeletal muscle. All experiments in the subsequent figures and supplemental figures were conducted using mt-cpYFP transgenic line 18.

Cardiac myocytes from transgenic mt-cpYFP line 18 mice also exhibited a significantly lower mitoflash frequency than that in FDB fibers isolated from the same animals (Fig. 4A left), while mitoflash frequencies in mitochondria isolated from the heart and skeletal muscle and incubated in 10 mM succinate were not different (Fig. 4A right). Mitoflash amplitude was significantly larger in FDB fibers compared to that observed in cardiac myocytes (Fig. 4B left), but this difference was reversed in mitochondria isolated in 10 mM succinate (Fig. 4B right). Mitoflash duration was not significantly different between FDB fibers and cardiac myocytes, but was modestly reduced in cardiac mitochondria compared to that observed for mitochondria isolated from skeletal muscle (Fig. 4C).

Figure 4. Mitoflash activity in skeletal and cardiac muscle is regulated by cell-specific environment.

(A-C) Average (±SEM) mitoflash frequency (A), amplitude (B) and duration (C) in FDB and cardiac myocytes (left) or isolated skeletal and cardiac mitochondria (right). Dramatic differences in mitoflash frequency and amplitude are observed between intact cells and isolated mitochondria. *p<0.05 compared with corresponding skeletal muscle preparations, Student-t test. n=10 cells from 3 mice for FDB; n=12 cells from 3 mice for cardiac myocytes; n=28 measurements from 4 mice for isolated skeletal mito; n=21 measurements from 3 mice for cardiac mitochondria. (D-F) Pearson Correlation analysis between large mitoflash duration (FDHM) and amplitude events in isolated skeletal mitochondria (D), isolated cardiac mitochondria (E), and the combination of both datasets (F). n=42–49 events for D and E. n=91 for F.

Inverse correlation between mitoflash amplitude and duration in isolated mitochondria

Close examination of mitoflash amplitude and kinetics in isolated mitochondria revealed the presence of two populations: 1) mitoflashes with long duration and small amplitude (Supplemental Fig. 5A) and 2) mitoflashes with large amplitude and short duration (Supplemental Fig. 5B). This observation suggests that mitoflash amplitude and duration might be inversely correlated. To test this idea, we filtered all mitoflash events using a criterion of ~200% of the mean value for each parameter, to segregate the two types of events. Events that fell above this criterion for either FDHM or amplitude were pooled for each tissue and analyzed using linear regression (Fig. 4D–F). Interestingly, mitoflash events exhibiting either a long duration or a high amplitude showed a strong inverse correlation in mitochondria isolated from both skeletal muscle and the heart (R²=0.69 for skeletal mitochondria and R²=0.78 for cardiac mitochondria; Fig. 4D & E), while the majority of the “normal” events fell within the shaded area below each line. This strong inverse correlation between large duration and high amplitude events remained even when all events from skeletal and cardiac mitochondria were combined (Fig. 4F). However, mitochondria isolated from skeletal muscle tended to have more events with longer duration, while events in mitochondria isolated from the heart favored a higher amplitude. This finding provides an explanation for the relatively shorter FDHM and larger amplitude mitoflash events observed in isolated cardiac mitochondria (Fig. 4B & C, right panels).

Mitochondrial substrates differentially regulate mitoflash activity in intact skeletal and cardiac muscle cells

We previously reported that mitoflash frequency in mitochondria isolated from skeletal muscle was similar in the presence of either complex I substrate (10 mM glutamate and 5 mM malate or Glu/Mal) or complex II substrate (5 mM succinate) [11]. For mitochondria isolated from the heart, mitoflash frequency was also similar in the presence of either complex I substrate (5 mM Glu/Mal) or complex II substrate (10 mM succinate) (Supplemental Fig. 6A). In both isolated mitochondrial preparations, mitoflash amplitude was slightly reduced in the presence of complex I substrate (Supplemental Fig. 6B and [11]). A similar substrate regulation of mitoflash frequency was also observed in cardiac mitochondria isolated from CypD KO mice (7.8 ±0.9 flashes/1000 mitochondria/100 s versus 10.0±2.2 flashes/1000 mitochondria/100 s with 10 mM succinate and 5 mM Glu/Mal, respectively).

While the above observations indicate that mitoflash properties and substrate dependence are similar for mitochondria isolated from the heart and skeletal muscle, mitoflash frequency and amplitude were very different when monitored in intact cells (Fig. 5). Differential substrate availability/dependence provides a potential explanation for the tissue-specific mitoflash properties observed above in intact cells (Fig. 4A and B, left). FDB muscle, a fast twitch muscle, exhibits a robust glycolytic system while fatty acid ß-oxidation represents the primary energy source for the ETC and tricarboxylic acid (TCA) cycle in cardiac muscle. Therefore, we hypothesized that the difference in mitoflash frequency in FDB fibers and cardiac ventricular myocytes is due, at least in part, to differences in mitochondrial substrate availability in the heart and skeletal muscle. The mitoflash results in FDB fibers and cardiac myocytes in Fig. 4 were conducted in the presence of 10 mM glucose, which might not be an ideal substrate for cardiac myocytes that prefer to burn fat. Therefore, we first determined the effect of a common fatty acid substrate, palmitoyl-L-carnitine (PC), on mitoflash activity in isolated cardiac mitochondria. PC produced a dose-dependent increase in both mitoflash frequency and amplitude. Specifically, mitoflash frequency (Supplemental Fig. 6C) and amplitude (Supplemental Fig. 6D) were significantly increased in 60 μM PC compared to that observed for 10 μM PC. We then examined the response of FDB fibers and cardiac myocytes to incubation in either Na-Pyruvate (NaPy) or Na-Palmitate (NaPal, added with L-Carnitine as a transporter). Both 5 mM NaPy and 50 μM NaPal+200 μM L-Carnitine caused only a modest (20–30%) increase in mitoflash frequency in FDB fibers compared with in the presence of 10 mM glucose (Glu) (Fig. 5A). In contrast, application of NaPy and NaPal enhanced mitoflash activity in intact cardiac myocytes ~3 fold (Fig. 5B). On the other hand, differences in mitoflash amplitude between FDB fibers and cardiac myocytes remained largely unchanged across the different substrates (Fig. 5C). No differences in mitoflash duration were observed across all conditions in both FDB and cardiac myocytes. Finally, stimulation of mitoflash activity by NaPy was similarly observed in FDB fibers and cardiac myocytes from CypD KO mice (Fig. 6A & B). Application of CsA in FDB fibers did not alter NaPy stimulation of mitoflash activity. Mitoflash amplitude and duration in FDB fibers and cardiac myocytes from CypD KO mice showed similar tissue-specific differences to those observed in WT cells (Fig. 5C & D for WT, Fig. 6C & D for CypD KO), except that mitoflash duration in CypD-deficient cardiac myocytes was slightly longer in NaPy compared to that in glucose (Fig. 6D, grey bars). In accordance with our observation that mitoflash frequencies were unchanged in FDB fibers and cardiac myocytes from CypD KO mice compared with that observed in WT cells in the presence of 10 mM glucose (Fig. 1D), mitoflash frequencies in FDB fibers and cardiac myocytes from CypD KO mice in the presence of NaPy (Fig. 6A & B) were also not significantly different from those observed for WT mice in the presence of NaPy (Fig. 5A & B).

Figure 5. Na-Pyruvate and Na-Palmitate application induced a considerably larger increase in mitoflash frequency in cardiac myocytes than that in FDB fibers from WT mice.

(A) Average (±SEM) mitoflash frequency in FDB fibers in the presence of 10 mM glucose (Glu), 5mM Na-Pyruvate (NaPy) or 50 μM Na-Palmitate (NaPal) plus 200 μM L-carnitine. *p<0.05 compared with Glu, one-way ANOVA with Tukey post-hoc test; n=32–47 cells from 4–5 mice. (B) Same as A, but using cardiac myocytes. Inset: same data replotted on an expanded scale. *p<0.05 compared with Glu, one-way ANOVA with Tukey post-hoc test; n=7–17 cells from 3 mice. (C-D) Average (±SEM) mitoflash amplitude (C) and duration (D) in FDB fibers and cardiac myocytes in the presence of 10 mM Glu, 5mM NaPy or 50 μM NaPal plus 200 μM L-carnitine (from the same set of experiments shown in A and B).*p<0.05 compared with FDB fibers, Student-t test; # p<0.05 compared with Glu one-way ANOVA with Tukey post-hoc test.

Figure 6. Na-Pyruvate induced a larger increase in mitoflash frequency in cardiac myocytes than that in FDB fibers from CypD KO mice or in the presence of CsA.

(A) Average (±SEM) mitoflash frequency in FDB fibers in the presence of 10 mM glucose (Glu), 5 mM Na-Pyruvate (NaPy), 10 mM glucose with CsA or 5 mM Na-Pyruvate with CsA. *p<0.05 compared with Glu, one-way ANOVA with Tukey post-hoc test; n=8–22 cells from 2–3 mice. (B) Average (±SEM) mitoflash frequency in cardiac myocytes in the presence of 10 mM glucose (Glu) or 5 mM Na-Pyruvate (NaPy). Inset: same data replotted on an expanded scale. *p<0.05 compared with Glu, Student-t test; n=12–17 cells from 2–3 mice. (C-D) Average (±SEM) mitoflash amplitude (C) and duration (D) in FDB fibers and cardiac myocytes from CypD KO mice in the presence of 10 mM glucose (Glu), 5 mM Na-Pyruvate (NaPy), 10 mM glucose with CsA and 5 mM Na-Pyruvate with CsA (from the same set of experiments shown in A and B for each tissue type).*p<0.05 compared with FDB fibers, # p<0.05 compared with Glu Student-t test.

DISCUSSION

The objective of this study was to investigate the mechanisms that underlie tissue-specific regulation of mitoflash activity by CypD and available mitochondrial substrates. In contrast to that reported previously [1–3, 22], our results do not support a tissue-specific regulation of basal mitoflash activity by CypD in skeletal and cardiac muscle. Specifically, we found that mitoflash properties, including frequency, amplitude and duration, were similar in both FDB fibers and cardiac myocytes isolated from WT and CypD KO mice (Fig. 1). This result was further confirmed in isolated mitochondria from skeletal muscles and hearts of WT and CypD KO mice (Fig. 2). These results suggest that CypD is not required for mitoflash generation under basal conditions in skeletal and cardiac muscle, at least under the conditions used in this study.

The reason for the difference between our findings and the conclusions of previous studies [1–3, 22] is unclear, although differences in genetic background may play a role. The mice used in the current study are on two mixed genetic backgrounds (C57B6N/SJL for mt-cpYFP and C57Bl6/129 for CypD KO mice), while mice used in other studies were on a congenic C57Bl6J background [1, 22]. In our studies, basal mitoflash frequency is relatively low in intact cardiac myocytes incubated in 10 mM glucose (1.3–1.8 flashes/1000 μm²/100 s) compared to that reported previously in cardiac myocytes under similar conditions [1, 25] or in langendorff perfused hearts [22]. Thus, the lower level of cardiac mt-pYFP expression in our transgenic lines could limit detection of low amplitude events in the current study. Furthermore, a limitation related to event detection due to low basal frequency (i.e. ability to resolve a reduction from an already low basal rate) is unlikely to be the case as a similar lack of CypD-dependence in mitoflash frequency in cardiomyocytes was observed in the presence of NaPy, where mitoflash frequency is 3 times higher than that in the presence of glucose (compare NaPy in Fig. 5B and Fig. 6B). Shang, et al attributed the lack of CypD-dependent regulation of mitoflash activity in skeletal muscle to a lower CypD expression. Thus, another possibility for the lack of CypD-dependence of mitoflashes in cardiomyocytes observed here is an adaptation in our compound mt-cpYFP/CypDKO mice that results in a reduction in CypD expression in the heart. This possibility could be examined by a quantitative comparison of CypD expression in the heart across all of the different mouse models used in mitoflash studies.

Consistent with prior studies [1, 22], we found that acute exposure to 10 mM CATR similarly increased mitoflash activity in both myocytes and isolated mitochondria from hearts of WT and CypD KO mice (Fig. 2F and Supplemental Fig. 3B and D). An increase in mitoflash frequency by acute exposure to CATR was similarly observed in FDB fibers (Supplemental Fig. 3C). We previously reported that pre-treatment for 20 mins with 1 μM CATR reduced mitoflash frequency in FDB fibers [3]. The difference between the two studies is likely due to CATR pretreatment inducing apoptosis through the activation of mPTP [26, 27], which would be less likely to occur during acute exposure. This result also suggests that CATR exerts its effect directly on the adenosine nucleotide translocator (ANT) to activate mPTP-independent of CypD.

While we show that CypD is not required for mitoflash generation in skeletal and cardiac muscle under basal conditions, CypD-dependent regulation of mPTP opening and mitoflash activity does have important physiological and pathophysiological implications. For example, our findings indicate that CypD-meditated mPTP function regulates activity-dependent changes in mitoflash frequency in skeletal muscle. Specifically, we found that CypD-deficiency results in a greater increase in mitoflash activity in FDB fibers following a series of high-frequency tetanic stimuli, consistent with CypD limiting activity-dependent augmentation of mitoflash generation (Fig. 3). As mitoflash frequency is enhanced by increases in mitochondrial Ca²⁺ in FDB fibers and cardiac myocytes [3, 15, 16] and CypD-deficient FDB fibers exhibit increased mitochondrial Ca²⁺ uptake following repetitive tetanic stimulation (Fig. 3E & F), CypD appears to regulate mitoflash activity by facilitating Ca²⁺ efflux from the mitochondria through Ca²⁺-dependent opening of the mPTP. This action may serve to limit mitochondrial Ca²⁺ overload and excessive ROS production during intense muscle activity. Consistent with this idea, stimulation-dependent increases in mitoflash activity were also abolished by pre-incubation of FDB fibers with Ru360, an inhibitor of mitochondrial Ca²⁺ uptake (Fig. 3D).

CypD-mediated regulation of mPTP opening and mitoflash activity have important implications for several pathological conditions including ischemia reperfusion [28], mitochondrial myopathy [29, 30], Ullrich Congenital Muscular Dystrophy/Bethlem Myopathy [31, 32] and amyotrophic lateral sclerosis (ALS) [33, 34]. A recent study reported that mitoflash activity is increased during a pre-symptomatic stage (2 months) in ALS mice (hSOD1G93A) and is further exacerbated after disease onset (4 months), when an elevation in CypD expression is observed [35]. Moreover, transient expressing hSOD1G93A in FDB fibers of WT mice results in a similar increase in mitoflash activity and CypD overexpression and application of CsA restores mitoflash activity to control levels [35]. In addition, an increase in mitoflash activity is also observed in denervated skeletal muscle fibers, which is abolished by application of CsA [36]. Therefore, while not being required for mitoflash generation, CypD is an important regulator of mitoflash activity under both physiological and pathophysiological conditions.

Karam et. al. [36] reported that a single 350 ms stimulation at a moderate frequency (40Hz) induced a significant and persistent (>120s) reduction in mitoflash activity in both WT and denervated FDB fibers, in contrast to the increase in mitoflash frequency following five 500 ms high frequency (100Hz) stimulation observed in our studies (Fig. 3 and [3]). While the studies used different stimulation protocols, this is unlikely to completely account for the differences in mitoflash regulation observed between these studies. The studies also used different stimulation approaches (local stimulation vs. field stimulation) and ages of mice (4 months vs 2.5–3 months). In addition, the studies also used different methods to determine mitoflash frequency. Specifically, our studies calculate the number of flashes occurred during the acquisition period as an indicator for flash frequency, while flash area (active area) was used in Karam et al. It is also interesting to note that representative images for mitoflashes shown in Karam et al. are mostly large patch flashes. Small individual flashes, rather than large patch flashes, are typically and most commonly observed in our studies. This observation suggests that there are some fundamental differences in the skeletal muscle fibers used in the two studies. However, one thing that is clear is that our stimulation protocol does not produce significant fiber damage since the increase in mitoflash frequency after stimulation recovers to the baseline levels <3 minutes after stimulation. In addition, the same repetitive tetanic stimulation protocol was also used to assess activity-dependent mitochondrial Ca²⁺ uptake in which fibers are monitored continuously 10 minutes after stimulation with no change in morphology and basal levels of myoplasmic Ca²⁺.

While the precise mechanisms that control mitoflash biogenesis remain incompletely elucidated, the events involved in determining mitoflash termination are even less understood. One hypothesis is that ROS levels during a mitoflash reach a critical threshold that activates a signal (e.g. depolarization of the mitochondrial membrane potential) that serves as a “safety valve” to terminate the event. Our finding of an inverse correlation between mitoflash amplitude and duration (Fig. 4D–E) support this idea. That is, mitoflash duration is short when its amplitude is large, consistent with a threshold for termination being reached rapidly for large amplitude events. Candidate targets for the “safety valve” function include any large conductance channel that is activated by ROS or pH, such as the mPTP, uncoupling proteins (UCPs) or the inner membrane anion channel (IMAC). Since the mPTP exhibits multiple sub-conductance states, it is possible that different mPTP conductances could account for the inverse relationship between mitoflash amplitude and duration. While overexpression of uncoupling protein type 2 (UCP2; the cardiac UCP isoform) reduces mitoflash frequency in cardiac myocytes, the effect on mitoflash duration was not reported [25, 37]. We recently found that neither mitoflash frequency, amplitude, nor duration were significantly changed in FDB fibers isolated from UCP3 KO mice (the skeletal muscle UCP isoform) transiently expressing mt-cpYFP [38]. Nevertheless, further studies in intact cells and isolated mitochondria are needed to determine the role of UCP proteins in mitoflash termination.

We found that mitoflash frequency in isolated mitochondria from skeletal and cardiac muscle is similarly dependent on mitochondrial substrates, although some differences in mitoflash amplitude and duration were observed. Therefore, the dramatic difference in mitoflash frequency between intact FDB fibers and cardiac myocytes is largely cell environment-specific. Direct application of TCA cycle substrate NaPy and fatty acid NaPal to facilitate β oxidation increased mitoflash frequency 3-fold in cardiac myocytes (Fig. 5), consistent with previous studies [17, 22]. The much smaller response in mitoflash frequency of FDB fibers to NaPyr and NaPal indicates that substrate availability/usage is at least in part responsible for the difference in basal mitoflash activity in skeletal and cardiac muscle. These results are consistent with cardiac myocytes exhibiting lower glycolytic efficiency than FDB fibers, which is not surprising given that cardiac muscle is a more oxidative muscle and FDB muscle is composed primarily of fast glycolytic muscle fibers. As mitoflash frequency of cardiac myocytes in the presence of NaPy and NaPal is still considerably lower than that of FDB fibers, difference in additional cellular factors and/or regulatory mechanisms (e.g. mitochondrial cardiolipin content) likely play a role in limiting mitoflash activity in the heart and/or enhancing mitoflash activity in skeletal muscle. Our observation that mitoflash activity was similarly regulated by NaPy in the absence of CypD is consistent with the CypD-independence of mitoflash activity in FDB fibers and cardiac myocytes.

In summary, this study investigated the relative role of CypD and mitochondrial substrates in regulating mitoflash activity in skeletal and cardiac muscle. While having no effect on basal mitoflash activity in either intact cells or isolated mitochondria, CypD limited activity-dependent increases in mitoflash frequency following repetitive, tetanic stimulation in a manner that depended on mitochondria Ca²⁺ uptake. We also showed that mitoflash amplitude and duration are inversely correlated, consistent with a “safety valve” mechanism for mitoflash termination. Finally, mitoflash activity in skeletal and cardiac muscle was strongly dependent on cellular context, with differences in mitochondrial substrate availability/usage being an important factor in differences in basal mitoflash activity between intact skeletal and cardiac muscle cells.

Supplementary Material

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Supplemental movie 1: Representative 60s x-y time series of mitoflash events from cardiac mitochondria isolated from a WT mouse (measurement in Fig. 2C left). The movie was accelerated 15× from real time. Circles indicate several representative mitoflash events. Scale bar: 20μm.

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Supplemental movie 2: Representative 60s x-y time series of mitoflash events from cardiac mitochondria isolated from a CypD KO mouse (measurement in Fig. 2C right). The movie was accelerated 15× from real time. Circles indicate several representative mitoflash events. Scale bar: 20μm.