Inhibition of MCU forces extramitochondrial adaptations governing physiological and pathological stress responses in heart

Significance

Mitochondrial Ca²⁺ is a fundamental signal that allows for adaptation to physiological stress but a liability during ischemia-reperfusion injury in heart. On one hand, mitochondrial Ca²⁺ entry coordinates energy supply and demand in myocardium by increasing the activity of matrix dehydrogenases to augment ATP production by oxidative phosphorylation. On the other hand, inhibiting mitochondrial Ca²⁺ overload is promulgated as a therapeutic approach to preserve myocardial tissue following ischemia-reperfusion injury. We developed a new mouse model of myocardial-targeted transgenic dominant-negative mitochondrial Ca²⁺ uniporter (MCU) expression to test consequences of chronic loss of MCU-mediated Ca²⁺ entry in heart. Here we show that MCU inhibition has unanticipated consequences on extramitochondrial pathways affecting oxygen utilization, cytoplasmic Ca²⁺ homeostasis, physiologic responses to stress, and pathologic responses to ischemia-reperfusion injury.

Abstract

Myocardial mitochondrial Ca²⁺ entry enables physiological stress responses but in excess promotes injury and death. However, tissue-specific in vivo systems for testing the role of mitochondrial Ca²⁺ are lacking. We developed a mouse model with myocardial delimited transgenic expression of a dominant negative (DN) form of the mitochondrial Ca²⁺ uniporter (MCU). DN-MCU mice lack MCU-mediated mitochondrial Ca²⁺ entry in myocardium, but, surprisingly, isolated perfused hearts exhibited higher O₂ consumption rates (OCR) and impaired pacing induced mechanical performance compared with wild-type (WT) littermate controls. In contrast, OCR in DN-MCU–permeabilized myocardial fibers or isolated mitochondria in low Ca²⁺ were not increased compared with WT, suggesting that DN-MCU expression increased OCR by enhanced energetic demands related to extramitochondrial Ca²⁺ homeostasis. Consistent with this, we found that DN-MCU ventricular cardiomyocytes exhibited elevated cytoplasmic [Ca²⁺] that was partially reversed by ATP dialysis, suggesting that metabolic defects arising from loss of MCU function impaired physiological intracellular Ca²⁺ homeostasis. Mitochondrial Ca²⁺ overload is thought to dissipate the inner mitochondrial membrane potential (ΔΨm) and enhance formation of reactive oxygen species (ROS) as a consequence of ischemia-reperfusion injury. Our data show that DN-MCU hearts had preserved ΔΨm and reduced ROS during ischemia reperfusion but were not protected from myocardial death compared with WT. Taken together, our findings show that chronic myocardial MCU inhibition leads to previously unanticipated compensatory changes that affect cytoplasmic Ca²⁺ homeostasis, reprogram transcription, increase OCR, reduce performance, and prevent anticipated therapeutic responses to ischemia-reperfusion injury.

Entry of Ca²⁺ into the mitochondrial matrix is a central event for Ca²⁺ homeostasis in cardiomyocytes (1) as well as for coordinating fundamental and diverse responses to physiological (2) and pathological stress (3). The paradigm for Ca²⁺ as a physiological second messenger that enhances oxidative phosphorylation to enable fight-or-flight responses but in excess contributes to disease and dysfunction is well established in myocardium (4). The molecular identity of the mitochondrial Ca²⁺ uniporter (MCU) was recently discovered, enabling development of new genetic models to understand the role of MCU in vivo. MCU is an ion channel protein that acts as the primary pathway for Ca²⁺ entry into the mitochondrial matrix (5, 6). Recent findings in global Mcu^−/− mice (7) suggest that the MCU pathway is dispensable for regulating cellular energy production, except under extreme physiological stress, and for activation of pathways leading to cell death; however, the effect of selective myocardial MCU inhibition is unknown. We developed a new transgenic mouse model with myocardial delimited dominant negative (DN)-MCU protein overexpression to test the role of MCU-mediated Ca²⁺ entry for myocardial physiology and pathological stress.

We tested whether loss of MCU-mediated Ca²⁺ entry substantially alters myocardial energetics. Surprisingly, we found that DN-MCU hearts had a higher oxygen consumption rate (OCR) due, at least in part, to secondary actions on cytoplasmic Ca²⁺ homeostasis. We also found that chronic MCU inhibition failed to protect against myocardial ischemia-reperfusion injury despite reducing generation of reactive oxygen species (ROS). We queried mRNA expression in adult hearts and identified diverse changes in multiple gene pathways induced by DN-MCU expression. Our findings reveal in vivo physiologic and pathological roles for cardiac MCU and suggest that loss of mitochondrial Ca²⁺ entry increases OCR, elevates cytoplasmic Ca²⁺, and sensitizes extramitochondrial cell death pathways.

Results

Increased Myocardial Oxygen Consumption and Reduced Performance in DN-MCU Hearts.

MCU, the pore-forming subunit of the mitochondrial Ca²⁺ uniporter, consists of two transmembrane domains that span the inner mitochondrial membrane and a linker-loop sequence (5, 6). The highly conserved aspartic acid-isoluecine-methionine-glutamic acid motif contains two negatively charged amino acids in the pore-forming linker-loop sequence. DN-MCU with D261Q/E264Q mutations inhibited mitochondrial Ca²⁺ uptake in HeLa cells (6). Based on this information, we developed a myocardial-selective in vivo model of MCU inhibition by transgenic expression of DN-MCU under control of the α-myosin heavy chain (αMHC) promoter (8) (Fig. 1A). DN-MCU mice were interbred into a CD1 background, based on evidence that CD1 background is permissive for loss of MCU current (9). DN-MCU mice were born in Mendelian ratios and survived into adulthood. We used a primer set to detect MCU and DN-MCU transcripts and found that the transcript level of Mcu was 60-fold higher in transgenic samples (Fig. 1B). The Myc-tagged DN-MCU protein was resident only in cardiac mitochondria from DN-MCU transgenic mice (Fig. 1C) and was detectable with an MCU antibody that showed markedly increased expression in DN-MCU compared with WT heart lysates (Fig. S1).

Fig. 1.

Increased myocardial oxygen consumption in DN-MCU hearts. (A) Schematic of the DN-MCU construct and expressed mutant channel in the transgenic mice. (B) Quantitative PCR measurement for WT and DN Mcu transcript expression. (C, Top) Western blot detection of MYC-tagged protein in heart (H), liver (L), and skeletal muscle (S) tissues from DN-MCU and WT mice. GAPDH was used as a loading control. (Bottom) Western blot detection of MYC-tagged protein in mitochondria (M) or cytosolic (C) isolates from DN-MCU and WT hearts. GAPDH confirmed purity of cytosolic isolates, and COXIV confirmed purity of mitochondrial isolates. (D) Oxygen consumption rates in Langendorff-perfused and paced hearts (beats/min). (E) Representative isolated hearts. (Scale bar, 200 μm.) (F) Summary of heart weight (HW) to body weight (BW) ratio measurements. (G) Left ventricular ejection fraction measured by echocardiography in unanesthetized mice. (H) Representative transmission electron microscopy images (5,000× and 10,000×). (I) Summary data of mitochondrial injury scores. (J) Mitochondrial protein (mg) measurements normalized to heart weight (g). (K) Mitochondrial:nuclear DNA. All error bars represent SEM. *P < 0.05, **P < 0.01, Student’s t test. Sample size (n) indicated for each group in parentheses.

Fig. S1.

MCU protein is increased in DN-MCU hearts. Western blot detecting MCU in mitochondrial isolates from DN-MCU and WT hearts. Coomassie stain was used as a loading control.

Increased mitochondrial Ca²⁺ can enhance oxidative phosphorylation (10). Based on the known relationship between mitochondrial Ca²⁺ and oxidative phosphorylation, we initially hypothesized that DN-MCU hearts lacking Ca²⁺ entry through MCU would have reduced O₂ consumption rates (OCR) compared with WT. Contrary to our expectations, unloaded Langendorff-perfused and ventricular paced DN-MCU hearts consumed more O₂ at 400 (P < 0.05), 600 (P < 0.01), and 750 beats/min (bpm) (P < 0.01) compared with WT (Fig. 1D). OCR was increased in WT between 400 and 600 bpm (P < 0.01) but not between 600 and 750 bpm. OCR was increased in DN-MCU between 400 and 600 bpm (P < 0.0001) as well as between 600 and 750 bpm (P < 0.05). No differences in cardiac morphology or baseline heart rate (Fig. 1E and Fig. S2) or in the heart weight:body weight ratios (Fig. 1F) were observed. We measured left ventricular (LV) ejection fraction in conscious, unsedated mice and found no difference between groups (Fig. 1G). No differences were detected between groups in the mitochondrial injury score (3) (Fig. 1 H and I), total mitochondrial protein content normalized to heart weight ratios (Fig. 1J), or mitochondrial-to-nuclear DNA content (Fig. 1K). Additionally, cyclooxygenase 4 (COXIV) protein levels were not different between groups (Fig. S3). These findings suggest that the increase in OCR in DN-MCU hearts was not related to alterations in myocardial or mitochondrial mass or structure but that DN-MCU hearts were less efficient than WT, based on higher OCR.

Fig. S2.

Echocardiographic analysis. Heart rate (HR), left ventricular (LV) mass, stroke volume, end systolic volume (ESV), and end diastolic volume (EDV), as measured by echocardiography. All error bars represent SEM. Sample size (n) is indicated for each group in parentheses.

Fig. S3.

COXIV protein expression is similar in WT and DN-MCU hearts. (Top) DN-MCU and WT whole-heart lysates were blotted for cyclooxygenase IV (COXIV) and Actin. (Bottom) The ratio of COXIV:Actin was used to compare mitochondrial protein content to cytosolic protein. Error bars represent SEM. Sample size (n) is indicated for each group in parentheses. No significant differences were detected.

DN-MCU Expression Decreases Inotropic and Lusitropic Responses to Stress.

In vivo LV pressure measurements showed that DN-MCU mice had increased baseline +dP/dt_MAX (the maximal LV pressure change rate during systole) and similar −dP/dt_MAX compared with WT. DN-MCU mice showed reduced ±dP/dt_MAX responses to isoproterenol (10 μg/kg) compared with WT (Fig. S4 A–F). Based on the defect in myocardial performance in DN-MCU mice in vivo, we repeated the Langendorff-perfused heart studies under conditions suitable for measuring LV pressure. We found that DN-MCU and WT hearts had equal LV-developed pressure (LVDP) at 400 bpm, but DN-MCU hearts had significantly reduced LVDP at 600 and 750 bpm compared with WT (P < 0.01) (Fig. 2 A–G). DN-MCU hearts had reduced LVDP between 400 and 750 bpm (P < 0.01) and 600 and 750 bpm (P < 0.05). +dP/dt_MAX and –dP/dt_MAX were not different at 400 bpm, but DN-MCU hearts had diminished ±dP/dt_MAX responses at 600 bpm (P < 0.01) and 750 bpm (P < 0.01) (Fig. 2 G and H). The +dP/dt_MAX was reduced in DN-MCU hearts between 400 and 750 bpm (P < 0.01) and 600 and 750 bpm (P < 0.05) and had significantly diminished –dP/dt_MAX between 400 and 750 bpm (P < 0.05) (Fig. 2I). Under these conditions DN-MCU hearts had significantly higher OCR at 400 and 600 bpm (P < 0.05) (Fig. 2J). Within groups, OCR at 400 and 750 bpm was significantly different (P < 0.01) in WT, but not in DN-MCU, suggesting that DN-MCU hearts contracting against an afterload have a smaller pacing-induced OCR range than unloaded hearts.

Fig. S4.

In vivo left ventricular pressure recordings. (A) Baseline +dP/dt_MAX (mmHg/s). (B) +dP/dt_MAX (mmHg/s) 2 min after isoproterenol administration (10 μg/kg, i.p.). (C) Percentage increase in +dP/dt_MAX from baseline to isoproterenol condition. (D) Baseline −dP/dt_MAX (mmHg/s). (E) −dP/dt_MAX (mmHg/s) 2 min after isoproterenol administration (10 μg/kg, i.p.). (F) Percentage decrease in −dP/dt_MAX from baseline to isoproterenol condition. (G) Baseline heart rate (HR) (bpm) measured by pressure waveform. (H) HR 2 min after isoproterenol administration (10 μg/kg, i.p.). (I) Percentage increase in HR from baseline to isoproterenol condition. All error bars represent SEM. Sample size (n) is indicated for each group in parentheses. *P < 0.05, ***P < 0.001, Student’s t-test.

Fig. 2.

DN-MCU expression reduced left ventricular pressure responses to pacing. (A and B) Representative waveform of left ventricular pressure (LVP) (mmHg) in WT and DN-MCU at 400 bpm, respectively. LVDP indicated by arrows. (C and D) Representative LVP changes in WT and DN-MCU hearts when increasing pacing rate from 400 to 600 bpm. (E and F) Representative LVP changes in WT and DN-MCU hearts when increasing pacing rate from 600 to 750 bpm. (G) LVDP (mmHg) at 400, 600, and 750 bpm. (H) +dP/dt_MAX (mmHg/s) at 400, 600, and 700 bpm. (I) −dP/dt_MAX (mmHg/s) at 400, 600, and 700 bpm. (J) OCR [μmol/min/g_(Wet)] in WT and DN-MCU hearts. All error bars represent SEM. Sample size (n) indicated for each group in parentheses. *P < 0.05, **P < 0.01, Student’s t test. #P < 0.05, ##P < 0.01, and ###P < 0.001 comparing 750 to 400 bpm; $P < 0.05 comparing 750 to 600 bpm, Tukey’s post hoc multiple comparison test.

DN-MCU Expression Alters Mitochondrial and Cytoplasmic Ca²⁺ Dynamics.

We next tested whether DN-MCU expression in ventricular myocytes prevented rapid mitochondrial Ca²⁺ entry. We used freshly isolated adult ventricular myocytes with permeabilized cell membranes and incubated them with Ca²⁺ green-5N (CaGr5N), a membrane-impermeable Ca²⁺ sensitive fluorescent dye (3). We confirmed that DN-MCU ventricular myocytes and isolated mitochondria had complete or nearly complete loss of mitochondrial Ca²⁺ uptake (Fig. 3A and Fig. S5A), similar to phenotypes observed in cells treated with the MCU antagonist Ru360 (11), lacking MCU expression (7) or expressing DN-MCU (12).

Fig. 3.

DN-MCU expression alters cytoplasmic Ca²⁺ dynamics. (A) Normalized kinetic tracings for CaGr5N-loaded, cell membrane-permeabilized ventricular myocytes. Arrows represent addition of 100 μM Ca²⁺. (B) Western blot detection of phosphorylated (pPDH) and total pyruvate dehydrogenase (PDH). (C) Summary data for the pPDH to total PDH ratio. (D) Summary data for PDH activity normalized to mitochondrial protein. (E) Representative Ca²⁺ transient traces from WT (black) and DN-MCU (red) ventricular myocytes stimulated by field stimulation. Summary data for (F) diastolic and (G) systolic [Ca²⁺] measurements made with Fura-2–loaded cells stimulated by field stimulation. All error bars represent SEM. *P < 0.05, ***P < 0.001, Student’s t test. Sample size (n) indicated for each group in parentheses.

Fig. S5.

Mitochondrial Ca²⁺ uptake measurement. (A) Assessment of Ca²⁺ uptake by isolated mitochondria using mitochondrial impermeant dye Calcium Green-5N. (B) Assessment of Ca²⁺ uptake by isolated mitochondria preloaded with intramitochondrial calcium indicator Fura-FF-AM.

To determine if mitochondrial Ca²⁺ ([Ca²⁺]_mt) was different between DN-MCU and WT hearts, we tested [Ca²⁺]_mt in isolated mitochondria loaded with Fura-4FF. We found that DN-MCU mitochondria had no observable step-wise increase in Fura-4FF signal in response to repetitive Ca²⁺ boluses (Fig. S5B). Furthermore, Ca²⁺ uptake in mitochondria isolated from DN-MCU hearts was unaffected by the MCU antagonist Ru360 (Fig. S5B), confirming that DN-MCU expression ablated the Ru360-sensitive Ca²⁺ influx. Taken together, these data show that DN-MCU myocardial mitochondria lack the MCU-mediated, rapid Ca²⁺ uptake pathway.

Mitochondrial Ca²⁺ stimulates pyruvate dehydrogenase phosphatase to dephosphorylate pyruvate dehydrogenase (PDH), increasing its enzymatic activity (13). We measured PDH phosphorylation in DN-MCU and WT hearts and found that PDH was significantly (P < 0.001) more phosphorylated (Fig. 3 B and C) and exhibited lower enzyme activity (Fig. 3D) in DN-MCU compared with WT cardiac mitochondria. To test whether glucose metabolism was altered in DN-MCU hearts, we used a Langendorff perfusion model with buffer containing [1,2-¹³C₂]-glucose followed by NMR analysis. The ¹³C incorporation was not different between groups (Fig. S6A), and aspartate was the only significantly increased metabolite in DN-MCU compared with WT hearts (Fig. S6 B and C). These data indicate that the lack of MCU-mediated Ca²⁺ uptake in DN-MCU mitochondria is sufficient to impair activity of the Ca²⁺-sensitive enzyme PDH, but without widespread changes in myocardial glucose metabolism detectable by NMR.

Fig. S6.

DN-MCU expression does not alter glucose metabolism in the heart. (A) Total peak intensity (in arbitrary units), a measure of total ¹³C incorporation obtained by summation of the peak intensities present in the heteronuclear multiple quantum coherence (HMQC) spectra. (B) Measured NMR peak intensity (in arbitrary units) of glucose metabolites in the perfused heart extracts. The peak intensities were measured from the assigned HMQC spectra (**P < 0.01 compared with WT, Student’s t test). (C) Overlay of the HMQC spectra of glucose metabolites of WT (black) and DN-MCU (red) perfused heart extracts. Select cross-peaks are labeled: lactate (Lac), alanine (Ala), glutamate (Glu), aspartate (Asp), and phosphocreatine (PCr). The one-dimensional slices through the Asp C3 are shown to display significant differences in peak intensity between the WT and DN-MCU heart extracts.

We next asked whether loss of mitochondrial Ca²⁺ uptake would increase cytosolic [Ca²⁺] in DN-MCU cells, potentially imposing an energy demand on the sarcoplasmic reticulum/endoplasmic reticulum Ca²⁺-ATPase (SERCA2a). We found significantly higher diastolic and systolic cytoplasmic [Ca²⁺] in DN-MCU cells (Fig. 3 E–G) compared with WT, suggesting that chronic MCU inhibition triggers a demand for increased SERCA activity. These findings show that chronic loss of MCU-mediated mitochondrial Ca²⁺ uptake in myocardium increased OCR, possibly by enhancing the metabolic cost of cytoplasmic Ca²⁺ homeostasis during excitation-contraction coupling.

We recently reported that cardiac pacemaker cells isolated from DN-MCU mice have impaired ATP production and defective cytosolic [Ca²⁺] homeostasis that was corrected by ATP dialysis (12). Based on these findings, we asked if ATP deficiency contributed to pacing-induced increases in cytoplasmic [Ca²⁺] in DN-MCU ventricular myocytes. Fortifying intracellular ATP (5 mM added to the pipette solution) significantly decreased cytosolic diastolic [Ca²⁺] only in DN-MCU ventricular myocytes (Fig. S7 A and B). In contrast, added ATP equally and slightly decreased systolic cytosolic [Ca²⁺] in WT and DN-MCU, but did not reach statistical significance (Fig. S7C). Taken together with findings in DN-MCU pacemaker cells (12), we interpreted these data to suggest that physiologic cytoplasmic [Ca²⁺] homeostasis requires MCU-mediated ATP production.

Fig. S7.

Reversal of elevated diastolic cytosolic [Ca²⁺] in voltage clamp-stimulated DN-MCU ventricular myocytes by ATP dialysis. (A) Representative [Ca²⁺] transients from isolated ventricular WT (black tracings) and DN-MCU (red tracings) myocytes stimulated at 1 and 3 Hz in the absence or presence of ATP (5 mM), which was added to the pipette (intracellular) solution. Summary data for diastolic (B) and systolic (C) cytosolic [Ca²⁺] in WT and DN-MCU ventricular myocytes in the presence and absence of pipette solution with 5 mM ATP. Cell number in each group is indicated in parentheses: (B) 1 Hz WT (7), WT + ATP (8), DN-MCU (16), DN-MCU + ATP (11); 3 Hz WT (8), WT + ATP (7), DN-MCU (13), DN-MCU + ATP (9). (C) 1 Hz WT (6), WT + ATP (6), DN-MCU (9), DN-MCU + ATP (6); 3 Hz WT (8), WT + ATP (7), DN-MCU (9), DN-MCU + ATP (10). ***P < 0.001, one-way ANOVA analysis of means. Error bars show SEM.

Multiple ionic conductances may contribute to differences in cytoplasmic [Ca²⁺] homeostasis between DN-MCU and WT ventricular myocytes. Therefore, we measured voltage-gated L-type Ca²⁺ current (I_Ca), Na⁺/Ca²⁺ exchanger current (I_NCX), and sarcoplasmic reticulum Ca²⁺ content (Fig. S8). We found no difference in I_Ca (Fig. S8A), but I_NCX density was increased (Fig. S8B) in DN-MCU cells, suggesting that DN-MCU cells partially rely on NCX to compensate for loss of MCU function. Interestingly, DN-MCU myocytes had reduced SR Ca²⁺ content at baseline (P < 0.01) and after isoproterenol treatment (P < 0.0001) compared with WT (Fig. S8C). We interpreted these data to suggest that MCU-mediated ATP production contributes to SR Ca²⁺ loading in ventricular myocytes and that MCU inhibition is accompanied by increases in I_NCX.

Fig. S8.

I_Ca and I_NCX recordings and SR Ca²⁺ content. (A) I_Ca recording in ventricular myocytes. Sample size (n) noted in parentheses. Error bars represent SEM. (B) I_NCX recording in patched ventricular myocytes. *P < 0.05, **P < 0.01, ***P < 0.001, Student’s t test. Sample size (n) noted in parentheses. Error bars represent SEM. (C) Summary SR Ca²⁺ content data from isolated ventricular myocytes loaded with Fluo-4 AM at baseline before and after addition of isoproterenol (ISO) (100 nM). Maximum SR Ca²⁺ release was triggered by a caffeine spritz (SI Materials and Methods). Sample size (n) is indicated for each group. **P < 0.01 and ****P < 0.0001, Student’s t test. Error bars show SEM.

No Effect of DN-MCU on OCR in Isolated Mitochondria.

Mitochondria are an important source of O₂^•- in cardiomyocytes (14) and may induce mitochondrial uncoupling to increase oxygen consumption. Therefore, we used electron paramagnetic resonance (EPR) spin trapping with the cyclic nitrone 5,5-dimethyl-1-pyrroline-N-oxide (DMPO) (15) to measure O₂^•- levels from isolated mitochondria. We found a trend (P = 0.07) toward less DMPO-OH signal in DN-MCU mitochondria (Fig. 4 A and B), suggesting reduced O₂^•- production. Importantly, when antimycin A, a complex III inhibitor and agent known to increase one-electron reductions of O₂ to form O₂^•-, was included in the reaction mixture the signals from WT and DN-MCU mitochondria increased to similar peak values (Fig. 4B), indicating that DN-MCU and WT mitochondria have a similar capacity to produce O₂^•-. Addition of superoxide dismutase (SOD) completely quenched the DMPO-OH signal in both DN-MCU and WT mitochondria, indicating that DMPO-OH was reporting on O₂^•- levels with high fidelity. These EPR data demonstrated that loss of MCU-mediated Ca²⁺ entry may modestly reduce O₂^•- production in isolated mitochondria.

Fig. 4.

DN-MCU mitochondria have normal OCR. (A) Representative electron paramagnetic resonance spectra of the DMPO-OH spin adduct produced by isolated cardiac mitochondria. (B) Summary showing the signal intensity of DMPO-OH baseline (P = 0.07, Student’s t test) and after addition of antimycin A (1 μM) or SOD (100 U/0.5 mL). (C) OCR in permeabilized myocardial fibers with 156 nM Ca²⁺ and 10 mM succinate in different states V₀ (no secondary substrate), V_ADP (1 mM ADP), and V_Oligo (1 mM oligomycin). Units are pmol oxygen/min/mg of dry fiber weight. (*P < 0.05 from WT in same state, Student’s t test). (D) Bioluminescent quantification of ATP (*P < 0.05 Student’s t test). (E) Calculated ATP:O ratios in permeabilized fibers. (F) Baseline OCR from isolated mitochondria without added ADP. (G) OCR normalized to baseline after addition of ADP (4 mM), oligomycin (2.5 μg/mL), carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) (4 μM), and antimycin A (4 μM). All error bars represent SEM. Sample size (n) indicated for each group in parentheses.

Excessive mitochondrial Ca²⁺ entry promotes opening of the mitochondrial permeability transition pore, loss of ΔΨm (16), and release of mitochondrial reactive oxygen species (ROS) (17), which are consequences of ischemia-reperfusion injury that lead to myocardial death (18). We adapted our ischemia-reperfusion model to simultaneously measure ΔΨm and ROS in Langendorff-perfused hearts using confocal microscopy. The DN-MCU hearts maintained ΔΨm at baseline values during ischemic stress, whereas WT hearts showed a 19 ± 6% decrease in ΔΨm during ischemia compared with baseline (Fig. S9A), although the difference in ΔΨm between genotypes was not significantly different (P = 0.12). In the reperfusion interval, WT hearts showed increased ROS compared with DN-MCU hearts (Fig. S9B). Toward the end of the reperfusion interval the DN-MCU hearts had significantly reduced ROS compared with baseline (P < 0.05), whereas WT hearts did not show a decline in ROS relative to baseline (Fig. S9B). Additionally, two-way ANOVA revealed that changes in ROS during ischemia reperfusion were significantly different between DN-MCU and WT hearts (P < 0.05). We considered the possibility that reduced ROS in DN-MCU mitochondria could be related to increased reductive enzyme activity. To test this concept, we measured glutathione peroxidase, catalase, Cu/ZnSOD, and MnSOD in freshly isolated whole hearts and found decreased activity of MnSOD and total SOD activity (Fig. S9 C–G) in DN-MCU compared with WT, suggesting that the reduced ROS signal in DN-MCU hearts during ischemia reperfusion was not due to increased antioxidant enzymes. Based on an extensive body of work (16, 19, 20), we initially hypothesized that lower levels of ROS would result in myocardial protection. However, despite reduction in ROS, the DN-MCU hearts were not protected from cell death following ischemia-reperfusion injury (Fig. S9 H and I).

Fig. S9.

DN-MCU hearts are not protected from ischemia-reperfusion injury. (A) Percentage change in TMRM signal from baseline in heart tissue during ischemia reperfusion. Two-way ANOVA analysis shows that the genotype was not a significant source of variation. (B) Percentage change in DCF signal from baseline in heart tissue during ischemia reperfusion, (*P < 0.05, **P < 0.01) compared with WT at same time point, Student’s t test; (#P < 0.05) compared with baseline value within genotype, Dunnett’s multiple comparison post test; (P < 0.05) Two-way ANOVA testing genotype as a source of variation. (C) Glutathione peroxidase activity. (D) Catalase activity. (E) Copper/zinc SOD-specific activity. (F) Manganese SOD-specific activity. (G) Total SOD activity. Sample size (n) is indicated for each group in parentheses. *P < 0.05, Student’s t test. Error bars show SEM. (H) Representative TTC-stained heart sections after ischemia reperfusion. The black line outlines viable tissue. (I) Quantification of viable myocardium from TTC staining. All error bars represent SEM. Sample size (n) is indicated for each group in parentheses.

Given the increased OCR in DN-MCU hearts, we next asked whether intrinsic properties of DN-MCU mitochondria contribute to high O₂ consumption. First, we studied isolated permeabilized myocardial fibers (21) (bath [Ca²⁺] = 156 nM) to assess OCR. We found that, in contrast to myocardium with intact cell membranes, DN-MCU fibers consumed significantly less O₂ than WT controls under state 3 conditions (P < 0.05), but had similar OCR relative to WT under state 2 and 4 conditions (Fig. 4C). ATP concentrations (Fig. 4D) were significantly lower (P < 0.05) in DN-MCU compared with WT samples, suggesting blunted ATP production, higher ATP hydrolysis, or both. The ratio of ATP content:O₂ consumption was not significantly different between DN-MCU and WT fibers (Fig. 4E). These findings suggest that DN-MCU mitochondria have reduced oxidative capacity compared with WT controls and are not intrinsically uncoupled, at least in the setting of membrane permeabilization and low extramitochondrial Ca²⁺.

Next, we extended our findings in disrupted myocardial fibers by determining OCR in isolated mitochondria in a buffer with nominally absent Ca²⁺. At baseline, we found that isolated DN-MCU mitochondria had similar OCR to WT (Fig. 4F). DN-MCU and WT mitochondria responded similarly to ADP, oligomycin, FCCP, and antimycin A (Fig. 4G). Thus, the reduction in mitochondrial respiration in permeabilized DN-MCU fibers and isolated mitochondria in low [Ca²⁺] compared with isolated hearts suggests that extramitochondrial [Ca²⁺] contributes to the increased OCR in intact DN-MCU hearts.

Broad Transcriptional Reprogramming in DN-MCU Hearts.

Our findings showed that inhibition of MCU has important consequences for cytoplasmic Ca²⁺ homeostasis and physiological and pathological stress responses in heart. As a first step toward identifying potential transcriptional foundations for these effects, we profiled transcriptional changes induced by inhibition of MCU by performing microarray analysis to measure mRNA expression levels. We detected 636 genes with more than twofold expression change (false discovery rate < 0.05) in adult DN-MCU mice, relative to WT littermates (Fig. 5A and Table S1, GSE62049). Functional gene annotation clustering revealed that these genes are significantly enriched for a variety of biological processes, including acetylation, redox biochemistry, and endoplasmic nuclear signaling (Fig. 5B). We used qRT-PCR to measure mRNA targets from selected functional gene annotation terms and validated these targets with working primer sets (Fig. 5C). These data support a concept that MCU-mediated mitochondrial Ca²⁺ entry has the potential to regulate transcriptional programs controlling diverse cellular pathways in myocardium. Our ANOVA analysis of mRNA expression levels revealed B-cell lymphoma 2 (Bcl2) to be elevated in DN-MCU samples (P = 0.00036). Therefore, we queried other members of the Bcl2 family and found markedly increased expression of the Bcl2-associated X protein (Bax) transcript (up-regulated ninefold, P = 0.006, Student’s t test) and significantly (P < 0.01) elevated Bax protein expression in DN-MCU heart lysates (Fig. 5 D and E). Bax is a Bcl-2 family member implicated in mitochondrial-dependent and -independent cell death pathways (22, 23) and has been shown to impact Ca²⁺ homeostasis (23), suggesting that DN-MCU hearts may have increased susceptibility to Bax-mediated death during ischemia-reperfusion injury.

Fig. 5.

Broad transcriptional reprogramming in DN-MCU hearts. (A) Hierarchical clustering of 700 differentially expressed genes (WT, black; DN-MCU, gray). (B) Graph shows P values for 10 functional terms enriched in the DN-MCU gene set. (C) qRT-PCR data showing validation of selected functional annotation terms listed in B. Superscript numbers indicate the functional annotation cluster being queried from B. (D) Western blot detecting Bax protein in whole-heart homogenates. Coomassie-stained blot shows gel loading for each sample. (E) Quantification of Bax Western blot (*P < 0.05, **P < 0.01, Student’s t test). All error bars represent SEM. Sample size (n) indicated for each group in parentheses.

Discussion

We found that loss of MCU causes unanticipated cellular responses that increase OCR, disrupt cytoplasmic Ca²⁺ homeostasis, and trigger transcriptional reprogramming. The ability of mitochondria to buffer cytosolic Ca²⁺ is controversial (24–26). Our findings show that inhibition of MCU increases cytosolic [Ca²⁺], potentially consistent with a loss of mitochondrial Ca²⁺-buffering capacity and/or ATP deficiency. We recently found that MCU inhibition limited fight-or-flight heart rate increases by lowering ATP below a critical threshold and that dialysis of 4 mM ATP was sufficient to rescue fight-or-flight responses in cardiac pacemaking cells (12). Thus, it is possible that elevated [Ca²⁺] in DN-MCU ventricular myocytes is at least partly due to inadequate ATP for physiological Ca²⁺ buffering.

We observed an abrogation of OCR differences using low [Ca²⁺] buffers and when mitochondria were tested in buffers nominally lacking Ca²⁺, suggesting that elevation of cytosolic [Ca²⁺] in DN-MCU hearts contributed to elevated OCR in situ. An elevated or depressed heart rate could increase or decrease OCR (27), but we controlled for this by measuring OCR at equal pacing intervals. Our in vivo data showed that DN-MCU mice had similar heart rates to WT at baseline, but an inability to increase heart rate after isoproterenol administration, consistent with recent evidence that MCU is necessary for heart rate increases during physiological stress (12). Rapid pacing in DN-MCU hearts may increase cytosolic [Ca²⁺] to a greater extent than in WT hearts due to loss of mitochondrial Ca²⁺ buffering and/or inadequate ATP to sustain intracellular Ca²⁺ homeostasis.

Our NMR glucose metabolite measurements did not reveal major differences in metabolites between DN-MCU and WT hearts. However, we cannot exclude the possibility that ¹³C was lost as CO₂ through the tricarboxylic acid (TCA) cycle because we measured labeled metabolites in clamp-frozen hearts and not effluents (28). The amount of glucose uptake in paced hearts is low (29) and was below the limit of detection in our NMR studies. Thus, it is possible that DN-MCU and WT hearts had differences in glucose uptake that were undetected under these experimental conditions.

By selectively eliminating MCU activity in myocardium, our studies revealed an unanticipated feature of the interdependence of cytosolic [Ca²⁺] and oxidative phosphorylation. Elevation in cytosolic [Ca²⁺] was substantially a consequence of impaired Ca²⁺-sensitive metabolism in the mitochondrial matrix because total ATP was reduced in DN-MCU hearts, compared with WT littermate controls, and because addition of exogenous ATP through a patch pipette improved cytoplasmic [Ca²⁺] in DN-MCU cardiomyocytes. To quantify the role of mitochondrial Ca²⁺ buffering in sculpting cytosolic [Ca²⁺] independently of metabolic actions, it would be necessary to develop a model with matrix resident Ca²⁺-activated dehydrogenases engineered for Ca²⁺ insensitivity.

Our study suggests that chronic manipulation of MCU is not a viable strategy to protect cardiomyocytes from ischemia-reperfusion injury. Acute MCU inhibition has shown promise as a therapeutic target to protect against cell death (30). We only tested for cell death in cardiomyocytes and cannot exclude the possibility that chronic MCU inhibition in different cell types could protect from pathologic stimuli. Our findings suggest that further advances in understanding mitochondrial mechanisms governing cell survival and cellular responses to loss of MCU-mediated mitochondrial Ca²⁺ entry are required before developing therapies designed to prevent mitochondrial Ca²⁺ overload.

Materials and Methods

A complete description can be found in SI Materials and Methods.

Mice Lacking Functional Myocardial MCU.

DN-MCU transgenic mice were recently described (12) and generated by αMHC promoter-driven expression of cDNA encoding the dominant negative form of MCU.

In Vivo and ex Vivo Measurements.

Hemodynamic and LV pressure measurements were made in anesthetized mice with a 1-F Millar catheter and in isolated, Langendorff-perfused hearts in the presence and absence of a LV pressure transducer.

Mitochondrial and Cytoplasmic Ca²⁺, OCR, and ROS Generation.

Extramitochondrial Ca²⁺ uptake was measured in cell membrane-permeabilized ventricular myocytes with CaGr5N, intramitochondrial Ca²⁺ measured with Fura-FF and cytoplasmic Ca²⁺ measured with Fura 2. OCR was measured in ex vivo hearts, cell membrane-permeabilized muscle fibers, and isolated mitochondria using SeaHorse Biosciences extracullular flux analyzer. O₂^•- was measured as SOD quenchable signal from isolated mitochondrial using EPR.

SI Materials and Methods

Ca²⁺ Green-5N Measurements.

We measured mitochondrial Ca²⁺ uptake using permeabilized ventricular myocytes as previously described (3).

Transmission Electron Microscopy.

LV free-wall tissue was isolated and processed as previously described (3).

Echocardiography.

Unanesthetized mice were used for echocardiography. The anterior chest was shaved and warmed gel was applied. The mouse was grasped gently by the nape of the neck and positioned in the left lateral position. A 30-MHz linear array transducer was applied to the chest to obtain cardiac images. The transducer was coupled to a Vevo 2100 imager (FUJIFILM VisualSonics). Images of the short and long axis were obtained with a frame rate of ∼180–250 Hz. All image analysis was performed offline using Vevo 2100 analysis software (Version 1.5). Endocardial and epicardial borders were traced on the short axis view in diastole and systole. The left ventricular length was measured from endocardial and epicardial borders to the left ventricular outflow tract in systole and diastole. The biplane area-length method was then performed to calculate left ventricular mass and ejection fraction.

In Situ Tetramethylrhodamine Methyl Ester and DCF Measurements.

Excised hearts were perfused with tetramethylrhodamine methyl ester (TMRM, 2 μM) and CM-H2DCFDA (DCF, 10 μM, Molecular Probes) containing Kreb–Henseleit’s (KH) solution (in mM: 120 NaCl, 24 NaHCO₃, 11.1 glucose, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 0.42 NaH₂PO₄, 10 taurine, 5 creatine, oxygenated with 95% O₂ and 5% CO₂) at room temperature for 30 min via the retrograde Langendorff perfusion system. Hearts were later transferred to another Langendorff apparatus (37 °C) attached to the confocal microscope system after TMRM and DCF loading was completed. The heart was placed onto a recording chamber for in situ confocal imaging of TMRM and DCF from epicardial myocytes under sinus rhythm. To avoid motion artifacts during imaging, Blebbistatin (10 μM, Sigma) and 2, 3-butandion-monoxim, 10 mM (Sigma) were added to the perfusion solution. Imaging of TMRM or DCF was achieved by tandem excitation at 488 and 561 nm, and the emission was collected at 500–545 and >575 nm, respectively. The heart was allowed to stabilize (10 min) before taking control images; the perfusion solution was switched to a mock ischemia solution (in mM: 135 NaCl, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 0.33 NaH₂PO₄, 10 Hepes) without 95% O₂ and 5% CO₂ for 30 min. The confocal frame images were acquired with confocal microscope (LSM510, Carl Zeiss MicroImaging Inc.) at 0, 5, 10, 15, 20, 30, and 40 min after reperfusion with KH solution. The microscope was equipped with 63× (N.A. = 1.4) oil immersion lens. The optical pinhole was set to 1 airy disk (<1 μm axial resolution) during confocal imaging. Each image frame contains 202 × 202 μm² area of myocytes. Five images from different locations of each left ventricular free wall were acquired, and intensity values from each ventricle were averaged to represent the global TMRM and DCF fluorescence intensity of each ventricle.

qRT-PCR.

RNA was isolated using TRIzol reagent (Invitrogen) and purified using a RNeasy MinElute Cleanup Kit (Qiagen). RNA concentrations were measured using a Nanodrop 2000 Spectrophotometer (Thermo Scientific). iScript cDNA Synthesis Kit (Bio-Rad) was used to generate cDNA from RNA using oligo(dT) primers. Validated PrimePCR primers (Bio-Rad) were used for qPCR on a ViiA 7 Real-Time PCR system (Applied Biosystems). Transcript levels were quantified by the ΔΔCt method.

In Situ O₂ and LV Pressure Measurements.

Isolated hearts were retrograde-perfused at 90 mm Hg with Krebs–Henseleit buffer containing glucose (10 mM) bubbled with 95% O₂/5% CO₂ at 37 °C and pH 7.4 (31, 32). The atrioventricular node was mechanically dissociated and hearts were paced (Bloom Electrophysiology) using a platinum pacing catheter (NuMed) positioned in the right ventricular apex. LVP was measured by inserting a balloon into the LV chamber through the mitral valve. The balloon was filled with degassed water and was connected to an APT-300 pressure transducer (World Precision Instruments). The signals were acquired with Axon Clampex software (Molecular Devices). The left ventricular end diastolic (LVEDP) was adjusted to 4–10 mmHg. The following parameters were measured: LV contractility parameters, the LVDP (the pressure difference between the peak LV systolic pressure and LVEDP) and the maximal LV pressure change rate during systole (+dP/dt_MAX), and the lusitropic parameter-minimum LV pressure change rate during diastole (−dP/dt_MAX). Cardiac O₂ consumption was calculated based on Henry’s Law as the difference between the O₂ partial tension measured in the perfusate prior (PO_2pre) and after (PO_2post) heart passage (Model NeoFox Instech Laboratories). The rate of O₂ consumption (VO_s) was calculated as: VO₂ = (PO_2pre – PO_2post)⋅α⋅v/m_heart, where v is the rate of coronary flow (T402, Transonic System Inc.), α = 0.024 mL O₂ per mL H₂O/760 mm Hg denotes the Brunsen’s solubility coefficient (31, 33), and m_heart is the wet heart weight. After 15 min of stabilization, only hearts with coronary flow rate between 2.5 and 5.0 mL⋅s⁻¹ and with LVDP greater than 80 mmHg were included for experimentation. The pacing protocol for O₂ consumption and LVP calculations was 9 min of continuous pacing consisting of 3 min at each of the following sequential cycle lengths: 150, 100, and 80 ms.

Permeabilized Fiber O₂ and ATP Measurements.

Mitochondrial function was studied using the saponin-permeabilized fiber technique (21) on heart muscle. State 2 respiration was measured after addition of 10 mM succinate. State 3 respiration was assessed by adding 1 mM ADP, and state 4 respiration was measured after addition of 1 mM oligomycin. Mitochondrial ATP measurements were performed by bioluminescence, as described (21).

Mitochondrial O₂ Measurements.

Hearts were excised and finely minced at 4 °C in a mitochondrial isolation buffer (70 mM sucrose, 210 mM mannitol, 5 mM Hepes, 1 mM EGTA, 0.5% BSA, pH 7.2). The minced heart was dounce-homogenized and then transferred to chilled Eppendorf tubes. Tubes were centrifuged at 4 °C twice at 600 × g, and supernatant was collected. A final spin of 10,000 × g at 4 °C was performed. The pellet was resuspended in mitochondrial assay buffer (70 mM sucrose, 210 mM mannitol, 10 mM KH₂PO₄, 5 mM MgCl₂, 2 mM Hepes, 1 mM EGTA, 0.2% BSA, 10 mM succinic acid, 2 μM rotenone, pH 7.2). Mitochondria were plated on a 96-well Seahorse Extracellular Flux Analyzer plate (Seahorse Biosciences) at a density of 0.5 μg/well. The plate centrifuged at 770 × g at 4 °C to ensure mitochondria stably present in the bottom of each well. The mitochondria assay buffer was replaced with prewarmed assay buffer. The addition of ADP (4 mM), oligomycin (2.5 μg/mL), FCCP (4 μM), and antimycin A (4 μM) were added sequentially with OCR measurements taken between each substrate addition.

Western Blotting.

Twenty micrograms of mitochondrial or heart lysate was loaded per well. Samples were in 4× LDS buffer (Life Technologies) containing RIPA, phosphatase (p0044, Sigma-Aldrich), and protease (Roche) inhibitors, and sample reducing agent (Life Technologies). Samples were run on 4–12% SDS/PAGE gels at room temperature and transferred to PVDF membrane overnight at 4 °C. pPDH (1:1,000, Cell Signaling), total PDH (1:1,000, Cell Signaling), MYC (1:1,000, Rockland), Bax (1:1,000, Cell Signaling), and MCU (1:500, Yenzyme) were incubated at room temperature for 2 h in TBST containing 5% milk. HRP-conjugated secondary antibodies were incubated at room temperature for 1 h.

Fura-2 Ca²⁺ Measurements.

Cytosolic Ca²⁺ levels were recorded from Fura-2–loaded cells, excited at wavelengths of 340 and 380 nm and imaged with a 510-nm long-pass filter. Single isolated ventricular myocytes were loaded with 4 µM Fura-2 acetoxymethyl (AM) for 30 min and then perfused for 30 min to de-esterfy the Fura-2 AM in normal Tyrode’s solution. After placement on a recording chamber, the cells were perfused in normal Tyrode’s solution at 33 ± 0.5 °C. Myocytes were stimulated with 1-, 3-, and 5-Hz bipolar pulse from SIU-102 (Warner Instrument).

L-Type Ca²⁺ Current Density Recordings.

As described (34), I_Ca was measured using whole-cell patch clamp technique at room temperature. I_Ca was confirmed by its sensitivity to Nifedipine 5 µM. Depolarizing voltage pulses (300 ms in duration) to various potentials (−70 to 60 mV in 10-mV steps) were applied from a holding potential of −80 mV. A prepulse to −40 mV of 50 ms was applied before each step to inactivate Na⁺ currents. The pipette (intracellular) solution comprised (mM): 120 CsCl, 10 Hepes, 20 tetraethylammonium (TEA) chloride, 1.0 MgCl₂, 0.05 CaCl₂, 10 glucose; pH was adjusted to 7.2 with 1.0 N CsOH. The bath (extracellular) solution comprised (mM): 137 NaCl, 10 Hepes, 10 glucose, 1.8 CaCl₂, 0.5 MgCl₂, and 25 CsCl; pH was adjusted to 7.4 with NaOH.

Na⁺/Ca²⁺ Exchanger Current Density Recordings.

I_NCX was recorded using whole-cell patch clamp technique at room temperature. The external solution contained the following (mM): 140 NaCl, 1.0 MgCl₂, 5.0 Hepes, 2.5 CaCl₂, 1.0 BaCl₂, 10 glucose, and 10 µM Nitrendipine, and 10 µM Strophanthidin (pH 7.4, adjusted with NaOH). The internal solution contained the following (mM): 110 CsCl, 10 NaCl, 5.0 glucose, 1.0 CaCl₂, 0.4 MgCl₂, 10 Hepes, and 20 TEA chloride, 5.0 EGTA (pH 7.2, adjusted with CsOH). Membrane currents were elicited by a command ramp of 2 s from +80 mV to −120 mV. Holding potential was −80 mV. The protocol was applied every 10 s. Recordings were taken when five consecutive ramp recordings showed nearly identical currents. I_NCX was measured as the Ni-sensitive current. NiCl₂ (5 mM) was added to define the fraction of current from NCX (the difference currents between total currents and post-Ni²⁺ currents from the same cell).

Ca²⁺ Transients Recording with Voltage Clamp Pacing.

Cytosolic Ca²⁺ levels were recorded from Fura-2–loaded ventricular cells, excited at wavelengths of 340 and 380 nm, and imaged with a 510-nm long-pass filter. Single isolated ventricular myocytes were loaded with 4 μM Fura-2 AM for 30 min and then perfused for 30 min to de-esterfy the Fura-2 AM in normal Tyrode’s solution. After placement on a recording chamber, the cells were perfused in bath solution comprised (mM): 137 NaCl, 10 Hepes, 10 glucose, 1.8 CaCl₂, 0.5 MgCl₂, and 25 CsCl; pH was adjusted to 7.4 with NaOH, at 35 ± 0.5 °C. Whole-cell patch clamp technique was used to dialyze cells. The pipette (intracellular) solution comprised (mM): 120 CsCl, 10 Hepes, 20 TEA chloride, 1.0 MgCl₂, 0.05 CaCl₂, 10 glucose; the pH was adjusted to 7.2 with 1.0 N CsOH. Adenosine 5′-triphosphate disodium salt hydrate 5 mM was added to pipette solution when needed. Myocytes were stimulated at 1 and 3 Hz using voltage protocol holding at −80 mV and step to 0 mV for 100 ms from a prepulse of 50 ms at −40 mV.

NMR Spectroscopy.

Hearts from 61-d-old male mice were perfused with KH solution containing 10 mM [1,2-¹³C₂]-glucose for 15 min and paced at 600 bpm. The hearts were ground in a precooled mortar, extracted with 4% perchloric acid, and centrifuged at 47,800 × g for 20 min at 4 °C. The supernatant was then neutralized using 1 M KOH and centrifuged again at 13,000 × g for 10 min at 4 °C. The resulting supernatant was lyophilized and dissolved in 50 mM sodium phosphate, pH 7.4, in 100% D₂O for NMR studies. ¹H homonuclear 2D DQF-COSY, TOCSY, and ROESY experiments, ¹H/¹³C heteronuclear 2D HMQC, HMBC, HSQC, and H2BC experiments, and ¹²C-filtered/¹³C-selected ¹H/¹H 2D COSY and TOCSY experiments were collected on the heart extract samples using a Bruker Avance II 800 MHz NMR spectrometer equipped with a sensitive cryoprobe for metabolite assignments. One-dimensional ¹³C spectra and ¹H/¹³C 2D ultra-high-resolution HSQC spectra with ¹J_13C-13C resolution were also acquired on a Bruker Avance II 500 MHz NMR spectrometer for ¹³C isotopomer quantification. NMR spectra of the heart extract samples were recorded at 25 °C. The ¹H and ¹³C chemical shifts were referenced to 2,2-dimethyl-2-silapentane-5-sulfonate. All NMR spectra were processed using NMRPipe package (35) and analyzed using NMRView (36).

Ischemia-Reperfusion Studies.

Excised hearts were perfused through the aorta with an oxygenated KH solution for 10 min. Thereafter, 30 min of no-flow global ischemia was induced. Following ischemia, hearts were reperfused with KH solution for 50 min. Hearts were flash-frozen for 2,3,5-triphenyltetrazolium chloride (TTC) staining. Briefly, hearts were serially sectioned and stained with 1% TTC in phosphate-buffered solution at 37 °C. Sections were then placed in 10% neutral buffered formalin and imaged on a stereoscope. Viable myocardium was compared with total tissue surface area using Image J.

Electron Paramagnetic Resonance.

Mitochondria were isolated from whole hearts in an MSE buffer (110 mM mannitol, 20 mM sucrose, 2 mM Tris⋅HCl, 1 mM DPTA, pH 7.2) by fine chopping with a razor and homogenization in a 5-mL Teflon dounce homogenizer. Mitochondria were pelleted and resuspended in a mitochondrial respiratory buffer (100 mM K-Aspartate, 20 mM KCl, 10 mM Hepes, 5 mM pyruvate, 1 mM glutamate, 1 mM malate, 0.3% fatty-acid-free BSA) at a concentration of 1 μg/1 μL. Mitochondrial samples (500 μL with 100 μg total protein) containing 100 mM of the spin trap DMPO (Dojindo Labs) were loaded into a standard quartz TM-flat cell (Wilmad LabGlass) and centered in a 4103 TM cavity (Bruker Instruments). Antimycin A (Sigma) and bovine erythrocyte Cu/Zn superoxide dismutase (Sigma), when added, were at concentrations of 1 μM and 100 units, respectively, with total reaction volume of 500 μL. ESR spectra were obtained while at room temperature with a Bruker EMX spectrometer at a frequency of 9.78 GHz and a modulation frequency of 100 kHz. Seven additive scans (1,024 points per x-scan) were signal-averaged with the microwave power at 39.85 mW, modulation amplitude 1.0 G, time constant 81.9 ms, scanning 80G/41.9 s, with a receiver gain of 5.02 × 10⁵. Signal heights were measured using the second low-field peak of the four-line DMPO spin adduct and are reported as arbitrary signal heights.

PDH Enzyme Assay.

The dipstick assay was performed in accordance with kit ab109882 (AbCam) using isolated mitochondria. Optical density was measured using Image J.

Gene-Expression Profiling Analysis.

Gene expression profiles for mouse heart tissues were performed using the Affymetrix Mouse 430A_2 chip as described (37). Partek Genomics Suite (Partek GS) was used for microarray data analysis. Affymetrix raw fluorescence intensity was quantified and normalized using multiarray analysis. Neonatal and adult sample data were used to perform a one-way ANOVA to detect differentially expressed genes in adult mice, but neonatal samples were not considered further. Functional annotation clustering was performed using The Database for Annotation, Visualization and Integrated Discovery (NIH).

Mitochondrial:Nuclear DNA Content Measurement.

DNA was extracted from apical ventricular myocardium using a DNEasy Blood and Tissue Kit (Qiagen). Mitochondrial (CCCATTCCACTTCTGATTACC, ATGATAGTAGAGTTGAGTAGCG) and nuclear (GTACCCACCTGTCGTCC, GTCCACGAGACCAATGACTG) targeted primers were used with SYBR Green mastermix (Bio-Rad) on an ABI StepOne Plus (Applied Biosystems) to compare mitochondrial to nuclear DNA ratio with the ΔΔCt method, as previously described (38).

In Vivo dP/dt Measurements.

Mice were anesthetized with ketamine/xylazine (87.5/12.5 mg/kg, i.p.) and placed supine on a sterile heating pad to maintain body temperature. A small incision was made in the anterior cervical region, and dissection was performed to expose the carotid artery. A 1.4-F Millar catheter was inserted into the artery and advanced into the left ventricle. Pressure was continuously recorded. ±dP/dt_MAX were analyzed just before isoproterenol treatment and 2 min after treatment with isoproterenol (10 μg/kg, i.p.) (LabChart, ADInstruments). Heart rates were calculated based on the characteristic pressure waves of the left ventricle.

Fura-FF Mitochondrial Ca²⁺ Uptake Measurements.

Mitochondria were isolated as described (25). Briefly, ventricles were homogenized in isolation buffer containing 75 mM sucrose, 225 mM mannitol, 1 mM Hepes, 1 mM EGTA, pH7.4, with 0.1 mg/mL bacterial proteinase type XXIV (Sigma-Aldrich). After addition of fatty-acid-free BSA (Calbiochem) to a final concentration of 0.2% (wt/vol), the homogenates were centrifuged at 500 × g for 5 min, and the supernatant was then centrifuged at 10,000 × g for 5 min. The mitochondrial pellet was then washed once with isolation buffer with 0.2% fatty-acid-free BSA and once without BSA. The mitochondria were then resuspended in resuspension solution (RS) with 75 mM sucrose, 225 mM mannitol, 1 mM Hepes, 20 µM EGTA, pH7.4, to 10–20 mg mitochondrial protein/mL as determined by Bradford protein assay (Bio-Rad).

Ca²⁺ uptake assays were carried out on a BioTek Synergy MX plate reader. For calcium uptake assay with extramitochondrial calcium indicator Calcium Green-5N (Life Technologies), 120 µg of mitochondria was added to each well of the 96-well assay plates containing assay solution with 1 µM Calcium Green-5N. The assay solution consists of 137 mM KCl, 20 µM EGTA, 20 mM Hepes, 5 mM glutamic acid, 5 mM disodium malate, 2 mM of potassium phosphate, pH 7.15. Free [Ca²⁺] was calculated by WEBMAXC STANDARD (web.stanford.edu/∼cpatton/webmaxcS.htm). Addition of 15.0, 17.6, 20.3, 25.0, and 35.0 µM of total [Ca²⁺] corresponded to 0.5, 1.0, 2.0, 5.7, and 15.3 µM increase of free [Ca²⁺], respectively. For calcium uptake assay with intramitochondrial calcium indicator Fura-FF-AM (Life Technologies), the mitochondria were first loaded in RS containing 20 µM Fura-FF-AM at room temperature for 30 min. After loading, mitochondria were washed twice with RS and then resuspended in RS and used for assay. When indicated, 5 µM Ru360 (Calbiochem) was added.

Laser-Scanning Confocal Imaging of Single Ventricular Myocytes.

Mouse ventricular myocytes isolated from WT or DN-MCU mice were loaded with Fluo-4 AM (5 μM) for 30 min at room temperature. After 20 min of de-esterification, the cell were placed in a recording chamber and perfused with normal Tyrode’s solution (1.8 mM Ca²⁺) at 36 ± 1 °C (Temperature Controller, TC2BIP, Cell MicroControls). Confocal Ca²⁺ imaging was performed in line-scan mode with a laser-scanning confocal microscope (LSM 510, Carl Zeiss) equipped with a N.A. of 1.35, 63× lenses. Images of Ca²⁺ transients and caffeine-induced Ca²⁺ transients (SR Ca²⁺ contents) were acquired at a sampling rate of 1.93 ms per line along the longitudinal axis of the cells. Steady-state Ca²⁺ transients were achieved by a 30-s pacing at 1 Hz. SR Ca²⁺ content was determined by measuring the amplitude of Ca²⁺ release induced by local delivery of 20 mM caffeine at baseline and after addition of 100 nM isoproterenol. All digital images were processed with IDL 6.0 program (Research System Inc.).

Measurement of Antioxidant Enzymes.

Mice were fully anesthetized and then euthanized. Hearts were rapidly excised and washed in ice-cold PBS and then flash-frozen in liquid nitrogen.

Catalase activity assay.

Catalase activity was determined on whole-mouse hearts, homogenized in 50 mM potassium phosphate buffer (pH 7.8, with 1.34 mM diethylenetriaminepentaacetic acid). Activity was measured spectrophotometrically. The disappearance of 10 mM hydrogen peroxide (240 nm) was measured as described (39). The equation A_60s = A₀ e^-kt was used to fit the exponential decay of H₂O₂ at 240 nm where k = the rate constant for the catalase-dependent disappearance of H₂O_2. Activity was normalized per mg protein (40) and expressed as mk units/mg protein as described.

SOD activity assay.

SOD activity was determined on mouse heart homogenates (as above) using an indirect competitive inhibition assay (41). The inhibition of superoxide-mediated nitroblue tetrazolium reduction (from xanthine/xanthine oxidase) was measured spectrophotometrically at 560 nm with one unit defined as the amount of protein capable of causing 50% of maximal inhibition. In this assay, configuration 1 unit corresponds to ∼6–10 ng for pure CuZnSOD/mL. MnSOD activity was determined by inhibition of CuZnSOD activity with 5 mM sodium cyanide. Activity is expressed as units/mg protein as determined by the Lowry method as described (42).

Glutathione peroxidase activity assay.

Selenium-dependent glutathione peroxidase activity (predominantly GPx1) was measured spectrophotometrically using 0.25 mM H₂O₂ as substrate with the method of Lawrence and Burk (43). One unit of enzymatic activity was defined as the amount of protein needed to oxidize 1 μmol NADPH/min and expressed as milliunits/mg protein as described (40, 42).