Heart rate increases are a fundamental adaptation to physiological stress, while inappropriate heart rate increases are resistant to current therapies. However, the metabolic mechanisms driving heart rate acceleration in cardiac pacemaker cells remain incompletely understood. The mitochondrial calcium uniporter (MCU) facilitates calcium entry into the mitochondrial matrix to stimulate metabolism. We developed mice with myocardial MCU inhibition by transgenic expression of a dominant-negative (DN) MCU. Here, we show that DN-MCU mice had normal resting heart rates but were incapable of physiological fight or flight heart rate acceleration. We found that MCU function was essential for rapidly increasing mitochondrial calcium in pacemaker cells and that MCU-enhanced oxidative phoshorylation was required to accelerate reloading of an intracellular calcium compartment before each heartbeat. Our findings show that MCU is necessary for complete physiological heart rate acceleration and suggest that MCU inhibition could reduce inappropriate heart rate increases without affecting resting heart rate.
Catecholamine agonists trigger physiological fight or flight increases in heart rate but the metabolic pathway(s) supplying adenosine triphosphate (ATP) for increasing heart rate are incompletely understood1,2. Cardiac pacemaker cells drive heart rate acceleration, at least in part, by augmenting energy-dependent flux of Ca2+ through an intracellular, sarcoplasmic reticulum (SR), storage compartment3. SR Ca2+ release triggers pacemaker cell membrane depolarization, leading to action potential initiation that triggers each heart beat4. Mitochondrial Ca2+ entry through the mitochondrial Ca2+ uniporter (MCU) can stimulate increased ATP production by enhancing the activity of dehydrogenases in the mitochondrial matrix that supply NADH for electron transport5,6. The recent discovery of the gene encoding the MCU protein Ca2+ permeation pore7,8 allowed us to test the potential role of MCU-mediated mitochondrial Ca2+ entry as a pathway for increasing ATP production to fuel heart rate increases. We developed new tools and approaches for studying the metabolic role of the MCU in cardiac pacing, including surgical gene transfer to pacemaker cells and transgenic mice with myocardial and pacemaker cell targeted expression of a dominant-negative (DN) MCU with pore domain mutations that prevented rapid, MCU-mediated mitochondrial Ca2+ entry. Here, we show that Ca2+ entry through the MCU is essential for telegraphing enhanced metabolic demand to pacemaker cell mitochondria and promoting oxidative phosphorylation. We found that isoproterenol (ISO) stimulates oxidative phosphorylation by the MCU pathway in cardiac pacemaker cells to fuel the activity of the sarcoplasmic-endoplasmic reticulum Ca2+ ATPase (SERCA2a), which is required for reloading SR Ca2+ stores and sustaining fight or flight heart rate increases. Inhibition of mitochondrial Ca2+ entry prevented increased oxidative phosphorylation, enhanced SERCA2a activity and physiological rate responses in cardiac pacemaker cells exposed to ISO. Dialysis of cardiac pacemaker cells with exogenous ATP rescued the fight or flight response to ISO despite MCU inhibition but ATP dialysis was ineffective after SERCA2a inhibition, by expression of a super-inhibitory phospholamban (PLN) mutant or thapsigargin, identifying SERCA2a as a critical control point downstream of MCU for heart rate increases and a preferential sink for mitochondrially-sourced ATP. Isolated hearts from wild-type mice with pacemaker-targeted DN-MCU gene therapy were resistant to rate increases by ISO. We found selectively obtunded ISO triggered rate increases in isolated pacemaker cells, in excised Langendorff-perfused hearts from wild-type mice with DN-MCU pacemaker-targeted gene therapy and in vivo in DN-MCU transgenic mice. Furthermore, DN-MCU transgenic mice showed reduced heart rates in response to spontaneous activity compared with wild-type littermate controls. In contrast to the profound loss of heart rate acceleration by MCU inhibition, unstimulated heart rates and autonomous pacemaker cell action potential firing were unaffected by the loss of MCU-mediated mitochondrial Ca2+ entry. Our findings highlight a previously unrecognized subcellular mechanism for catecholamine-triggered heart rate increases and provide insight into a role for MCU-mediated mitochondrial Ca2+ entry as a metabolic second messenger required for the physiological fight or flight stress response9. These results define the MCU as an essential activator of a metabolic pathway for heart rate control and suggest that MCU inhibition in cardiac pacemaker cells has therapeutic potential to selectively prevent excessive heart rates.
The MCU mediates rate increases in pacemaker cells
Isolated cardiac sinoatrial nodal (SAN) pacemaker cells spontaneously generate action potentials under basal conditions, in the absence of catecholamine stimulation. The rate of action potential initiation is increased with ISO, a catecholamine β-adrenergic receptor agonist, in a concentration-dependent manner (Fig. 1a–d)10. We found that SAN cells dialysed with Ru360 (5 μM), an MCU antagonist, had significantly reduced action potential frequency increases to ISO (Fig. 1b,d) compared with SAN cells without Ru360 (Fig. 1a,d). The inhibitory effect of Ru360 on SAN cell action potential frequency responses to ISO was reversed by co-dialysis with ATP (4 mM), a concentration present in heart cells (Fig. 1c,d)11. Co-dialysis with 1 and 2 mM ATP pipette solutions was inadequate to rescue ISO-mediated rate increases while 8 mM ATP did not result in greater rate responses than 4 mM ATP (Supplementary Fig. 1a–c), suggesting fight or flight SAN responses operate between an [ATP] threshold >2 and </=4 mM. In contrast, neither Ru360 nor ATP had any affect on basal SAN cell rates and ATP dialysis had no affect on SAN rate responses to ISO in the absence of Ru360 (Supplementary Fig. 1a,b), indicating that basal heart rate was independent of an MCU pathway and that exogenous ATP did not affect Ru360-independent cellular processes important for accelerating heart rate. We next corroborated our results with Ru360 dialysis by infecting SAN cells with adenovirus encoding a DN-MCU, containing pore domain charge reversal mutations that prevent MCU-mediated mitochondrial Ca2+ entry (Fig. 1e–h)7,8. Both Ru360 and DN-MCU were similarly effective at preventing mitochondrial Ca2+ entry (Supplementary Fig. 2) and, like intracellular dialysis with Ru360, DN-MCU expression in SAN cells interfered with ISO-induced rate increases (Fig. 1f,h). The DN-MCU-dependent loss of the ISO rate response was rescued by co-dialysis with ATP (Fig. 1g,h), mirroring findings with MCU inhibition by Ru360 (Fig. 1c,d). Cultured SAN cells, with and without DN-MCU expression, had slower basal spontaneous rates than freshly isolated counterparts (baseline in Fig. 1d,h) but nevertheless exhibited significant rate increases over baseline in response to ISO (Fig. 1i,j). We considered the possibility that the mitochondrial Na+/Ca2+ exchanger, a MCU-independent pathway for mitochondrial Ca2+ efflux, could also contribute to SAN rate increases12 by testing SAN responses to CGP-37157 (1 μM), a Na+/Ca2+ exchanger antagonist. After CGP-37157, we measured a 1.2±6.0% increase in WT SAN cells (n=8) and 0.8±2.6% increase in DN-MCU SAN cells (n=3, P=0.88), suggesting that the mitochondrial Na+/Ca2+ exchanger did not affect basal heart rate. After ISO (1 μM), we measured a 79.8±5.8% (n=4) increase from baseline rate in WT SAN cells and 32.0±11.1% increase from baseline (n=4) rate in DN-MCU SAN cells (P<0.01), similar to ISO-mediated increases in the absence of CGP-37157 (Fig. 1i). These findings suggested that the mitochondrial Na+/Ca2+ exchanger was unlikely to participate in SAN rate responses, consistent with other recent findings13. The rescue of physiological responses to ISO after MCU inhibition by ATP dialysis suggested that MCU activity was an upstream event required to supply adequate ATP for increasing heart rate.
MCU inhibition impairs heart rate acceleration
The selective control of pacemaker cell rate increases by MCU inhibition suggested that the MCU could be a novel target for controlling heart rate increases without slowing resting heart rates. To test this concept in hearts, we used a targeted gene-painting approach to deliver adenovirus expressing DN-MCU or eGFP to the SAN in vivo14. One week after SAN gene painting, we verified SAN targeted gene expression (Fig. 2a,b) and measured rate responses to ISO in excised, Langendorff-perfused hearts (Fig. 2c–f). We found DN-MCU SAN gene painting significantly and selectively reduced heart rate increases to ISO without affecting spontaneous heart rates in the absence of ISO. We next developed transgenic mice with myocardial DN-MCU expression (Fig. 3a,b and Supplementary Fig. 3) to test the role of MCU in heart rate acceleration in vivo. The DN-MCU mice had hearts with normal chamber size and function (Fig. 3c), detectable SAN DN-MCU expression (Fig. 3d), reduced ATP content (Fig. 3e) and complete loss of mitochondrial Ca2+ uptake in myocardial cells (Fig. 3f,g). DN-MCU hearts showed increased Mcu mRNA, as expected, and reduced mRNA expression for auxiliary MCU regulatory proteins (Supplementary Fig. 4), possibly consistent with compensatory transcriptional reprogramming in response to the loss of functional MCU. The reduced fight or flight response in DN-MCU hearts was selective for catecholamine agonist stimulation because DN-MCU and wild-type SAN cells had similar rate responses to BayK 8644 (1 μM), a voltage-gated Ca2+ channel agonist capable of accelerating heart rate independent of ISO. In the presence of BayK 8644 DN-MCU SAN cells (n=5) exhibited 69.9±10.9% rate increases over baseline, similar to published responses in wild-type SAN cells15. During echocardiography measurements, lightly sedated DN-MCU mice exhibited resting heart rates similar to wild-type littermate controls (Fig. 3c). Lightly sedated and restrained DN-MCU and wild-type mice during cutaneous electrocardiogram (ECG) recording showed similar resting heart rates (Fig. 3h–j), but reduced ISO-stimulated heart rate increases (Fig. 3j,k). DN-MCU mice had significantly prolonged P-waves, PQ and PR intervals but similar QRS and QT interval durations compared with WT controls, suggesting that loss of MCU current slows atrial and atrioventricular conduction velocity without affecting conduction velocity in the distal conduction system or in ventricular myocardium (Supplementary Table 1). Finally, we measured heart rates in unrestrained and unsedated mice with surgically implanted ECG and activity telemeters. DN-MCU mice showed modest but significant reductions in basal heart rate compared with wild-type controls and these differences were enhanced by activity (Fig. 3l,m) and ISO (Fig. 3n,o). Taken together, these data showed heart rate responses to spontaneous activity and ISO required MCU, whereas basal heart rates were independent of MCU, indicating MCU inhibition could selectively prevent heart rate increases in vivo.
MCU enables rapid refilling of SR Ca2+ in pacemaker cells
Catecholamine stimulation increases heart rate by enhancing SAN cell membrane inward current and shortening the time between action potential firing16. Release of intracellular Ca2+ from the SR provides the electrochemical driving force for the cell membrane Na+/Ca2+ exchanger inward current (INCX) in SAN cells17. Elimination of SR Ca2+ release by the toxin ryanodine or SERCA2a inhibition by thapsigargin (Supplementary Fig. 5) significantly reduced the SAN response to ISO, findings that demonstrate the required connection between SR Ca2+ release and physiological SAN cell acceleration18. To further test the apparent connection between MCU activity and SR Ca2+ flux, we performed confocal line-scan measurements on SAN cells isolated from DN-MCU and wild-type control mice (Fig. 4). DN-MCU SAN cells had significantly fewer diastolic Ca2+ release events and reduced SR Ca2+ content after ISO stimulation compared to wild-type controls, suggesting that impaired fight or flight responses by MCU inhibition occurred because of reduced SR Ca2+ flux during physiological stress. SERCA2a activity requires ATP to pump cytoplasmic Ca2+ to the SR lumen19, so we measured the rate of decay of the cytoplasmic Ca2+ concentration ([Ca2+]cyto) using a fluorescent indicator (Fura 2 AM, 0.1 μM) to test if SERCA2a was a sink for ATP produced by an Ru360-sensitive process. ISO significantly increased the rate of decline in [Ca2+]cyto compared with SAN cells in control bath solution, reflecting enhanced activity of SERCA2a (Fig. 5a)20. Ru360 dialysis slowed the decline in [Ca2+]cyto after ISO (Fig. 5b), while co-dialysis of ATP (4 mM) with Ru360 restored the rate of [Ca2+]cyto decline to ISO-stimulated values present in the absence of Ru360 (Fig. 5c,d). However, Ru360 did not slow the decline in [Ca2+] cyto in the absence of ISO stimulation (Fig. 5e), consistent with the lack of effect of Ru360 on SERCA2a activity or basal SAN cell action potential frequency. In contrast to the effect of ATP dialysis on SAN cells exposed to Ru360, ATP dialysis did not significantly increase the rate of [Ca2+]cyto decline (Fig. 5f–h) or SAN action potential frequency (Fig. 5i) in SAN cells isolated from mice expressing a super-inhibitory mutant form of phospholamban (N27A)21 that constrains SERCA2a despite ISO stimulation. These results show that ISO increases SAN cell rates by actions that require MCU and SERCA2a. We considered the possibility that MCU inhibition was somehow affecting the ability of ISO to enhance phospholamban phosphorylation, which reduces the inhibitory actions of phospholamban on SERCA2a22. We found that atrial tissues from DN-MCU and wild-type littermates had similar increases in phospholamban phosphorylation after ISO (Supplementary Fig. 6), suggesting that MCU inhibition did not interfere with SERCA2a activity by actions on phospholamban nor did DN-MCU expression promiscuously affect downstream signalling actions of ISO. We interpreted the rescue of ISO responses by exogenous ATP after the elimination of MCU-mediated Ca2+ entry but not after SERCA2a inhibition to suggest that MCU contributes to ATP synthesis targeted for SERCA2a consumption during physiological stress.
MCU is not a global effector of pacemaker currents
Because heart rate is responsive to multiple ionic currents16, we next asked if the MCU pathway affected the ATP and 3′-5′-cyclic adenosine monophosphate (cAMP)-dependent cell membrane ion channel (HCN4) inward current (If)23 an ionic current known to participate in SAN cell automaticity. We found that If responses to ISO in isolated SAN cells were not reduced by Ru360 dialysis (Fig. 6a,b). The lack of effect of Ru360 on If suggested that Ru360 did not reduce cAMP nor ATP availability globally in SAN cells below a threshold necessary to increase If. To test this concept further, we measured the maximum diastolic cell membrane potential, which is primarily determined by activity of the Na+/K+ ATPase16. MCU inhibition by Ru360 dialysis or by transgenic expression of DN-MCU did not affect the maximum diastolic membrane potential in isolated SAN cells (Fig. 6c). We also measured CaV1 L-type Ca2+ current (ICa), an SAN cell membrane inward current enhanced by ISO through ATP-mediated phosphorylation24. Similar to our findings with If, ISO-induced ICa increases were not impaired by Ru360 (Fig. 6d,e). These findings were consistent with a model where selective loss of heart rate acceleration after ISO by Ru360 was primarily or exclusively related to actions on SERCA2a.
MCU is required for ISO to increase NADH
Mitochondrial Ca2+ entry increases oxidative phosphorylation by enhancing the activity of key mitochondrial dehydrogenases to provide NADH/NADPH-reducing equivalents required for ATP synthesis25. This mechanism is activated when cellular Ca2+ enters the inner mitochondrial membrane space from the cytosol through the MCU pathway26. We first asked whether mitochondrial Ca2+ entry was critical for oxidative phosphorylation-dependent ATP synthesis in SAN pacemaker cells. We infected cultured mouse SAN cells with adenovirus encoding mt-pericam27, a circularly permutated Ca2+-sensitive fluorescent protein, to measure mitochondrial Ca2+ concentration ([Ca2+]mito). The mt-pericam expression was localized to mitochondria in adenovirus infected SAN pacemaker cells, based on co-localization with MitoTracker Orange (Fig. 7a). ISO caused an increase in [Ca2+]mito (Fig. 7b) that was prevented by dialysis of Ru360 (Fig. 3b–d). Mitochondrial Ca2+ enhances ATP production by augmenting NADH, the primary electron donor for electron transport28. We next measured NADH fluorescence at baseline and after addition of ISO. ISO increased NADH fluorescence and this increase was prevented by Ru360 (Fig. 8a,b, Supplementary Fig. 7a,b) and in SAN cells with transgenic DN-MCU expression (Supplementary Fig. 7c,d), suggesting that ISO enhancement of NADH required MCU-mediated mitochondrial Ca2+ entry. These data confirm the MCU-dependence of coupling between [Ca2+]cyto and [Ca2+]mito in SAN cells and show that the MCU provides critical metabolic support for SERCA2a activity and SR Ca2+ loading. Together these data are consistent with a model where MCU-mediated enhancement of oxidative phosphorylation is a metabolic mechanism enabling the physiological fight or flight stress response in cardiac pacemaker cells (Fig. 8c).
Our data provide new mechanistic understanding into the fight or flight response to physiological stress by showing that heart rate increases rely on the MCU in cardiac pacemaker cells. In contrast, basal rates do require SERCA activity but are independent of MCU, suggesting that availability of MCU-independent ATP production is sufficient to sustain heart rates in the absence of extreme physiological stress. Oxidative phosphorylation is enhanced by [Ca2+]mito, which is required to generate ATP that fuels SERCA2a activity under extreme physiological stress. Our findings show that Ca2+ homeostatic mechanisms in pacemaker cells form the framework for fight or flight heart rate increases but do not exclude additional modulation of heart rate by other Ca2+ sensitive signals29,30 or by Ca2+ independent ionic currents16. The MCU metabolic pathway appears optimized to generate heart rate increases during episodes of high energy demand that are signalled by catecholamines. Our data are consistent with earlier work showing that mitochondrial Ca2+ is required to optimize refilling of intracellular Ca2+ stores by SERCA2a31 and where ATP dialysis (3 mM) recovered Ca2+ sequestration by intracellular stores after the addition of mitochondrial toxins32. Our findings provide insight into recent work showing MCU knockout selectively impairs high workload activity in striated muscle33. However, our data show that basal pacemaker cell activity is uncoupled from MCU-dependent ATP production. Because the relationship between MCU and SERCA2a in pacemaker cells appears purposed to selectively enable heart rate acceleration, future therapies targeting MCU or SERCA2a in pacemaker cells could provide a means to fine tune heart rates by preventing excessive heart rates without reducing resting heart rates.
All the experiments were carried out in accordance with the guidelines of Institutional Animal Care and Use Committee (PHS Animal Welfare Assurance, A3021-01).
SAN cell isolation and electrophysiological recordings
Isolation of single SAN cells from mice was performed according to previously published methods29,34 with minor modifications. Mice (6–8 weeks, females and males) were administered an intraperitoneal injection of avertin (20 μl g−1) and monitored until unresponsive. The heart was excised and placed into Tyrode’s solution (35 °C), consisting of (mM) 140.0 NaCl, 5.0 HEPES, 5.5 glucose, 5.4 KCl, 1.8 CaCl2 and 1.0 MgCl2. The pH was adjusted to 7.4 with NaOH. The SAN region, delimited by the crista terminalis, atrial septum and orifice of superior vena cava, was dissected free from the heart. The SAN was cut into smaller pieces, which were transferred and rinsed in a solution containing (mM) 140.0 NaCl, 5.0 HEPES, 5.5 glucose, 5.4 KCl, 0.2 CaCl2, 0.5 MgCl2, 1.2 KH2PO4, 50.0 taurine and 1.0 mg ml−1 bovine serum albumin, with pH adjusted to 7.4 using NaOH. SAN tissue pieces were digested in 5 ml of solution containing collagenase type I, elastase (Worthington) and protease type XIV (Sigma) for 20–30 min. The tissue was transferred to 10 ml of Kraft–Bruhe medium containing (mM) 100.0 potassium glutamate, 5.0 HEPES, 20.0 glucose, 25.0 KCl, 10.0 potassium aspartate, 2.0 MgSO4, 10.0 KH2PO4, 20.0 taurine, 5.0 creatine, 0.5 EGTA and 1.0 mg ml−1 bovine serum albumin, with pH adjusted to 7.2 using KOH. The tissue was agitated using a glass pipette for 10 min. The cells were stored at 4 °C and studied within 7 h.
SAN cells were placed in Tyrode’s solution at 36±0.5 °C. SAN cells were identified by their characteristic morphology (spindle or spider shape) and spontaneous activity in all single-cell experiments. SAN cells were also identified electrophysiologically by typical spontaneous action potentials with slow depolarizing phase 4 and the hyperpolarization-activated current (If) in electrophysiological experiments.
Spontaneous action potentials and If were recorded using the perforated (amphotericin B or β-escin) patch-clamp technique35 on single SAN cell at 36±0.5 °C in Tyrode’s solution (0.5 mM BaCl2 was added to the bath solution when recording If36,37). The pipette was filled with (mM) 130.0 potassium aspartate, 10.0 NaCl, 10.0 HEPES, 0.04 CaCl2, amphotericin B 240 μg ml−1 with pH adjusted to 7.2 with KOH. SAN cells with stable action potentials lasting at least 5 min were included in the experiments. β-escin 25 μM was substituted for amphotericin B to allow dialysis of Ru360 or ATP. When recording If, membrane potential was held at −35 mV, the voltage steps were applied for 5 s ranging from −125 mV to −45 mV in 10 mV increments or vice versa38. ICa was measured using the perforated patch technique at 36±0.5 °C as previously described29. ICa was confirmed by its sensitivity to nifedipine 5 μM. Depolarizing voltage pulses (300 ms in duration) to various potentials (−60 mV to 60 mV in 10 mV step) were applied after 50 ms at −40 mV to inactivate INa from a holding potential of −70 mV. The pipette solution comprised (mM): 120.0 CsCl, 10.0 EGTA, 10.0 HEPES, 10.0 tetraethylammonium chloride, 5.0 phosphocreatine, 3.0 CaCl2, 1.0 MgATP, 1.0 NaGTP and the pH was adjusted to 7.2 with 1.0 N CsOH. The bath (extracellular) solution comprised (mM): 137.0 NaCl, 10.0 HEPES, 10.0 glucose, 1.8 CaCl2, 0.5 MgCl2, 25.0 CsCl, pH was adjusted to 7.4 with NaOH.
Viral infection with 5mt-pericam/eGFP and DN-MCU
Freshly isolated mouse (6–8 weeks, females and males) SAN tissue was cut into small pieces and put into 35 mm tissue culture plates with DMEM. Fresh medium containing Ad-eGFP, Ad-5mt-pericam or Ad-DN-MCU was added to the plates at a multiplicity of infection of 100. Ad-5mt-pericam and Ad-DN-MCU were generated as follows: Ratiometric-Pericam-mt27 was first subcloned into pacAd5CMV-mcs-KN (University of Iowa Gene Transfer Vector Core) using HindIII and EcoRI. As the localization of this Pericam with a single Cox4 targeting sequence was not exclusive to mitochondria, four additional mitochondrial targeting sequences were added to the amino terminus. Two tandem Cox8a targeting repeat units from the pcDNA3-D4cpv vector39 were subcloned via HindIII into pacAd5CMV-ratiometric-Pericam-mt. Colonies were screened to identify clones that contained four tandem Cox8a inserts and confirmed via sequencing. For Ad-DN-MCU containing a carboxy (C)-terminal Myc tag, human MCU cDNA clone (GenBank accession code BC034235) was first obtained from the I.M.A.G.E consortium (ID: 5296557) and subcloned into pAd5CMVmcsIRESeGFP (University of Iowa Gene Transfer Vector Core, Iowa City, IA, USA) by PCR using Phusion DNA Polymerase (New England Biolabs) and the GeneArt Seamless Cloning and Assembly Kit (Life Technologies). PCR primers amplifying Myc-tagged MCU were: forward 5′- ATAAGCTTATGGCGGCCGCCGCAGGTAGATCG -3′, reverse 5′- CTACAGGTCTTCTTCGCTAATCAGTTTCTGTTCATCTTTTTCACCAATTTGTCGGAG -3′, and pAd5CMVmcsIRESeGFP: forward 5′- GAAGAAGACCTGTAGGATATCGAATTCCTGCAGCCC -3′, reverse 5′- GCCGCCATAAGCTTATCGATACCGTCGACCTC -3′. Dominant-negative mutations in MCU that inhibit Ca2+ conductance (D260Q, E263Q)7,8 were generated with Agilent’s QuikChange site-directed mutagenesis kit. Positive clones were confirmed by DNA sequencing. Adenoviruses expressing of 5mt-Pericam and DN-MCU-myc were generated by the University of Iowa Gene Transfer Vector Core. Expression of the recombinant ratio-5mt-pericam and DN-MCU in SAN cells was detected by GFP fluorescence. Recombinant adenovirus that expresses eGFP only (Ad-eGFP) was used as a control.
NADH and mitochondrial Ca2+ measurements
The autofluorescence of endogenous NADH, which derives primarily from mitochondria40,41, was measured as described41. In brief, NADH was excited at 350 nm (AT350/50X, Chroma) and fluorescence was recorded at 460 nm (ET460/50m and T400LP, Chroma). We normalized NADH level with FCCP as 0%, Rotenone-induced NADH change as 100%. The baseline level of NADH was 22%±3 (n=15) of the rotenone-induced maximal value.
Mt-pericam, a mitochondrial matrix-targeted, circularly-permuted green fluorescent protein fused to calmodulin and its target peptide M13 (ref. 27). Pericam emission at 535 nm due to excitation at 415 nm reports changes in Ca2+ (ref. 42). We used 5 mt-pericam to measure Ca2+mito in isolated, cultured SAN cells. The mitochondrial Ca2+ level was monitored in cells transiently expressing the 5 mt-pericam protein at excitation wavelengths of 415 nm, presented as 1−F/F0 (Ca2+mito) (refs 42, 43, 44), and the emission collected using a 535-nm band-pass filter.
Intracellular Ca2+ transients
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 SAN cells were loaded with 0.1 μM Fura-2 AM for 20 min, and then perfused for 20 min to de-esterify 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 36 °C±0.5. Spindle-shaped, spontaneously beating cells were chosen for the experiments. Action potential recording was performed simultaneously.
SAN gene painting
SAN painting was performed as previously described14,45. Briefly, Poloxamer 407 (Spectrum), trypsin (Sigma) and collagenase, type II (Worthington) mixture was made with 40% Poloxamer, 1% trypsin and 0.25% collagenase in PBS, and then added to an equal volume of recombinant adenovirus expressing cDNA for the gene(s) of interest (DN-MCU-IRES GFP vs eGFP) in solution. This mixture was liquid in consistency at 4 °C but gelled at 37 °C. Mice (6–7 weeks, females and males) were anaesthetized using ketamine/xylazine (87.5/12.5 mg kg−1, respectively), intubated and ventilated. The junction of the superior vena cava and right atrium was visualized through a small incision in the second intercostal space. The gel was applied to the posterior surface of the junction of the superior vena cava and right atrium with a fine brush. The intercostal muscles, pectoralis major and minor and the skin incision were closed using 6/0 silk and the mice were allowed to recover.
Ex vivo Langendorff-perfused heart rate measurements
ECG recording from Langendorff-perfused hearts was performed as described29. Briefly, excised hearts from 7 to 9-week female and male mice were rapidly mounted on a modified Langendorff apparatus (HSE-HA perfusion systems, Harvard Apparatus, Holliston, Mass) for retrograde aortic perfusion at a constant pressure of 80 mm Hg with oxygenated (95% O2, 5% CO2) Krebs–Henseleit solution consisting of (mM) 25.0 NaHCO3, 118.5 NaCl, 4.0 KCl, 1.2 MgSO4, 1.2 NaH2PO4, 1.5 CaCl2 and 11.2 glucose, with pH equilibrated to 7.4. Each perfused heart was immersed in a water-jacketed bath and was maintained at 36 °C. ECG measurements from the intact heart were continuously recorded with Ag+-AgCl electrodes, which were positioned around the heart in an approximate Einthoven configuration. After the heart was allowed to stabilize for 15 min, different concentrations of isoproterenol were added to the perfusate.
The SAN area was dissected from freshly isolated hearts for comparing transgenic DN-MCU mice with WT or from Langendorff perfused hearts for localizing SAN gene painted tissue. The SAN tissue was imbedded in OCT tissue freezing media and cooled over liquid nitrogen before being stored at −80 °C. 10 μM serial sections were taken through the tissue. All sections were slowly brought from −20 °C to room temperature in 4% paraformaldehyde. Sections were washed with ice cold PBS, permeabilized (0.1% Triton X-100, 0.1% sodium citrate in PBS) and blocked (3% fish gelatin, 2 mg ml−1 BSA, 0.1% Triton X-100 in PBS). HCN4 (Abcam) (1:200) and Myc epitope tag (Rockland) (1:200) antibodies were incubated with samples overnight at 4 °C. Samples were washed in blocking buffer and fluorophore conjugated secondary antibodies (1:500) were incubated overnight. Gene painted sections were treated with DAPI and imaged on an EVOS FL Auto microscope. DN-MCU transgenic samples and WT sections were images on a confocal microscope (Zeiss).
Calcium green mitochondrial Ca2+ uptake assays
We measured mitochondrial Ca2+ uptake using permeabilized HEK cells as previously described46,47. Briefly, cells were grown in DMEM, with 10% FBS and 1% of Penicillin/Streptomycin. At 80% confluence cells were infected with DN- MCU adenovirus at MOI 10. Cells were harvested after 24 h incubation and placed into a 96 well plate. Each well was loaded with 1 million cells in respiratory buffer containing 125.0 mM KCl, 2.0 mM K2HPO4, 20.0 mM HEPES, 5.0 mM glutamate, 5.0 mM malate, 0.005% saponin, and 5 μM thapsigargin. 5 μM Ca2+ was injected at 3 min intervals. Fluorescence was measured using a Tecan plate reader. Adult cardiac myocytes were isolated from 6- to 8-week-old wild-type or DN-MCU TG mice using a previously described isolation procedure46. A total of 50 nM Blebbistatin was included in the myocyte buffer to prevent cellular contraction. Fluorescence intensity was measured from 50,000 cardiac myocytes per well. Then, 100 μM Ca2+ was injected at 3-min intervals for myocyte experiments.
DN-MCU overexpressing mice
The inter-membrane D260IME263 amino-acid motif of human MCU was mutated to the dominant-negative (DN) form8, QIMQ, by replacing the nucleotide sequence, 5′- gacatcatggag -3′ with 5′- cagatcatgcag -3′ using site-directed mutagenesis (agilent.com). The resulting DN-MCU DNA product was amplified by PCR with Phusion DNA polymerase using a forward primer containing a SalI restriction site (5′- ACCAACGTCGACATGGCGGCCGCCGCAGGTAG -3′) and reverse primer containing a C-terminal myc epitope tag sequence and HindIII restriction site (5′- GAACGCAAGCTTCCTACAGGTCTTCTTCGCTAATCAGTTTCTGTTCATCTTTTTCACCAATTTGTCGGAG -3′). PCR products were digested and ligated into SalI and HindIII digested pBS-αMHC-script-hGH vector and positive clones confirmed by sequencing. Mouse embryonic stem cells were injected with the linearized DNA (digested with NotI) in the University of Iowa Transgenic Mouse Core Facility and implanted into pseudo-pregnant females to generate B6XSJL F1 mice. Insertion of the transgene into the mouse genome was confirmed by PCR analysis (not shown) using the forward primer, 5′- CCCACACCAGAAATGACAGACAGAT -3′ and reverse primer, 5′- AGAGGAGCAGCAGGAGCGATCTA -3′, producing a product of 200 bases. Mice were backcrossed to F4 generation or greater into the CD1 background. Transgenic and control mice of either gender were killed at the age of 2–3 months.
Western blots for detecting phospholamban
Heart lysates were prepared from flash-frozen mouse right atria from 6- to 8-week-old female and male mice. Mice were injected with ISO (0.4 mg kg−1) 10 min before harvesting the right atrium and western blotting was performed with a SDS–polyacrylamide gel electrophoresis (SDS–PAGE) electrophoresis system as described29. Briefly, 30 μg protein samples were size-fractionated on SDS–PAGE, and then transferred to polyvinylidene difluoride membranes. The membranes were probed with anti-pSer16-PLN (1:5,000), anti-pThr17-PLN (1:5,000) (from Badrilla Ltd., Leeds, UK) at room temperature for 4 h. Then membranes were incubated with Alexa-Fluor680-conjugated anti-mouse (Invitrogen Molecular Probes, Carlsbad, CA, USA) and/or IR800Dyeconjugated anti-rabbit fluorescent secondary antibodies (Rockland Immunochemicals, Gilbertsville, PA, USA) and scanned on an Odyssey infrared scanner (Li-Cor, Lincoln, NE, USA). Integrated densities of protein bands were measured using ImageJ Data Acquisition Software (National Institute of Health, Bethesda, MD, USA). Uncropped scans of all western blot images are available in the SI.
Western blots for detecting MYC-tagged DN-MCU
Heart, liver and skeletal muscle tissues were harvested and flash frozen. Samples were homogenized in RIPA buffer (10 mM Tris-Cl, pH8.0, 1 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate, 0.1% SDS, 140 mM NaCl), containing protease and phosphatase inhibitors with antioxidant. Twenty micrograms of protein were fractionated on NuPAGE (Invitrogen) 4–12% SDS–PAGE gels and transferred overnight to polyvinylidene difluoride membranes (Bio-Rad). Nonspecific binding was blocked with 10% w/v non-fat milk powder in TBS-T (50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 27 mM KCl and 0.25% Tween-20). The membranes were probed with anti-MYC (1:1,000) (Rockland Immunochemicals, Gilbertsville, PA, USA) at room temperature for 2 h. Then membranes were incubated with anti-rabbit HRP-conjugated secondary antibody (1:5,000). Protein bands were visualized using ECL reagent (Lumi-Light, Roche). Correct loading was confirmed with Coomassie staining of membranes. Uncropped scans of all western blot images are available in the SI.
Mouse surface electrocardiograms
Mouse surface electrocardiogram (ECG) tracings were acquired as described48. Before the ECG acquisition 6- to 8-week-old female and male mice were pre-anaesthetized with 2% isoflurane in 1l oxygen per minute (Isotec100 Series, Isoflurane Vaporizer, Harvard Apparatus, Holliston, USA). Mice were placed in a supine position on a heated ECG pad (Mousepad, THM 100, Indus Intruments, Webster, USA), and the limbs were attached to the pad electrodes using a tape to obtain ECG lead II. Anaesthesia was maintained via facemask by continuous isoflurane ventilation as described above. The body temperature was continuously monitored using a rectal probe and sustained within 36–37 °C. To obtain a baseline ECG, animals were allowed to rest for 5 min after being positioned on the pad. Thereafter isoproterenol (10 μg kg−1) was injected intraperitoneally. ECG acquisition was performed continuously using a multichannel amplifier and data acquisition system (Powerlab 16/30, AD Instruments, Colorado Springs, USA) converting the signal into digital for a further data analysis (Labchart Pro software, version 7, AD Instruments, Colorado Springs, USA).
Laser scanning confocal imaging of single SAN cells
Laser scanning confocal imaging of single SAN cells were performed as described49. Mouse SAN cells isolated from WT or DN-MCU mice were loaded with Fluo-4 AM (5 μM) for 20 min at room temperature. After 20 min of de-esterification, the cells were placed in a recording chamber and perfused with normal Tyrode solution (1.8 mM Ca2+) at 36±1 °C (Temperature Controller, TC2BIP, Cell MicroControls). Spindle-shaped, active spontaneously beating cells were chosen for the experiments. Confocal Ca2+ imaging was performed in line-scan mode with a laser scanning confocal microscope (LSM 510, Carl Zeiss) equipped with a numerical aperture (NA) 1.35, × 63 lenses. Images of spontaneously beating 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. SR Ca2+ content was determined by measuring the amplitude of Ca2+ release induced by local delivery of 20 mM caffeine. All digital images were processed with IDL 6.0 program (Research System Inc).
Surgical implantation of ECG telemeters was performed as described29. In brief, 8- to 10–week-old female and male mice were anaesthetized with ketamine/xylazine (87.5/12.5 mg kg−1), and an ECG transmitter (DSI model Ta10EA-F20) was implanted in the abdominal cavity. The leads were placed subcutaneously in a lead I configuration. Experiments were performed after a 5-day recovery period. ECG and activity were recorded for 72 h from undisturbed mice at a sampling rate of 1 kHz. Heart rate and activity were calculated from serial 1-min averages. On the fourth day, there was a 1-h baseline ECG recorded before ISO injection followed by an ISO injection and an additional 1-h recording epoch. On the fifth day, we repeated this protocol with another dose of ISO. The last dose of ISO was given on the sixth day. The heart rate in response to ISO was calculated from serial 10-s averages.
Transthoracic echocardiography was performed as previously described50. Unanesthetized, sedated mice were used for echocardiography. 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 Visual Sonics, Toronto, Canada). Images of the short and long axis were obtained with a frame rate of ~180–250 hertz. All image analysis was performed offline using Vevo 2100 analysis software (Version 1.5).
The 6- to 8-week-old littermate mice were killed. Atria were rapidly harvested and flash frozen in liquid nitrogen. Atria were homogenized, sonicated and centrifuged at 10,000g. The supernatants were collected and a luciferase assay (Invitrogen, ATP Determination Kit A22066) was used to detect ATP. Samples were measured in triplicate on a Femtomaster FB 12 luminometer (Zylux).
Quantitative RT 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 PCR primers (Bio-Rad) were used for qRT PCR on a StepOnePlus Real-Time PCR system (Applied Biosystems). Transcript levels were quantified by the ΔΔCt method.
Data are presented as mean±s.e.m., unless otherwise noted. Statistical analysis was performed either with one-way analysis of variance or an unpaired or paired Student’s t-test, as appropriate. The Holm–Sidak test was used for post hoc comparisons after analysis of variance. Analyses were performed with Sigmaplot or Sigmastat (Systat Software, Inc. San Jose, CA 95110, USA). The null hypothesis was rejected for a P<0.05.
How to cite this article: Wu, Y. et al. The mitochondrial uniporter controls fight or flight heart rate increases. Nat. Commun. 6:6081 doi: 10.1038/ncomms7081 (2015).
Cannon, W. B. The Wisdom of the Body 312W W Norton & Co. (1932).
Bers, D. M. Excitation-Contraction Coupling and Cardiac Contractile Force 2nd edn, 427Kluwer Academic Press (2001).
Sirenko, S. et al. Ca2+-dependent phosphorylation of Ca2+ cycling proteins generates robust rhythmic local Ca2+ releases in cardiac pacemaker cells. Sci. Signal 6, ra6 (2013).
Lakatta, E. G., Maltsev, V. A. & Vinogradova, T. M. A coupled SYSTEM of intracellular Ca2+ clocks and surface membrane voltage clocks controls the timekeeping mechanism of the heart's pacemaker. Circ. Res. 106, 659–673 (2010).
Jouaville, L. S., Pinton, P., Bastianutto, C., Rutter, G. A. & Rizzuto, R. Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic priming. Proc. Natl Acad. Sci. USA 96, 13807–13812 (1999).
Denton, R. M. Regulation of mitochondrial dehydrogenases by calcium ions. Biochim. Biophys. Acta 1787, 1309–1316 (2009).
De Stefani, D., Raffaello, A., Teardo, E., Szabo, I. & Rizzuto, R. A forty-kilodalton protein of the inner membrane is the mitochondrial calcium uniporter. Nature 476, 336–340 (2011).
Baughman, J. M. et al. Integrative genomics identifies MCU as an essential component of the mitochondrial calcium uniporter. Nature 476, 341–345 (2011).
Glancy, B. & Balaban, R. S. Role of mitochondrial Ca2+ in the regulation of cellular energetics. Biochemistry 51, 2959–2973 (2012).
Irisawa, H., Brown, H. F. & Giles, W. Cardiac pacemaking in the sinoatrial node. Physiol. Rev. 73, 197–227 (1993).
Gupta, A., Chacko, V. P. & Weiss, R. G. Abnormal energetics and ATP depletion in pressure-overload mouse hearts: in vivo high-energy phosphate concentration measures by noninvasive magnetic resonance. Am. J. Physiol. Heart Circ. Physiol. 297, H59–H64 (2009).
Boyman, L., Williams, G. S., Khananshvili, D., Sekler, I. & Lederer, W. J. NCLX: the mitochondrial sodium calcium exchanger. J. Mol. Cell. Cardiol. 59, 205–213 (2013).
Liu, T. et al. Inhibiting mitochondrial Na+/Ca2+ exchange prevents sudden death in a Guinea pig model of heart failure. Circ. Res. 115, 44–54 (2014).
Swaminathan, P. D. et al. Oxidized CaMKII causes cardiac sinus node dysfunction in mice. J. Clin. Invest. 121, 3277–3288 (2011).
Gao, Z. et al. Catecholamine-independent heart rate increases require Ca2+/calmodulin-dependent protein kinase II. Circ. Arrhythm. Electrophysiol. 4, 379–387 (2011).
Mangoni, M. E. & Nargeot, J. Genesis and regulation of the heart automaticity. Physiol. Rev. 88, 919–982 (2008).
Sanders, L., Rakovic, S., Lowe, M., Mattick, P. A. & Terrar, D. A. Fundamental importance of Na+-Ca2+ exchange for the pacemaking mechanism in guinea-pig sino-atrial node. J. Physiol. 571, 639–649 (2006).
Rigg, L., Heath, B. M., Cui, Y. & Terrar, D. A. Localization and functional significance of ryanodine receptors during β-adrenergic stimulation in guinea-pig sino-atrial node. Cardiovasc. Res. 48, 254–264 (2000).
Verjovski-Almeida, S. & Inesi, G. Fast-kinetic evidence for an activating effect of ATP on the Ca2+ transport of sarcoplasmic reticulum ATPase. J. Biol. Chem. 254, 18–21 (1979).
Cavagna, M., O’Donnell, J. M., Sumbilla, C., Inesi, G. & Klein, M. G. Exogenous Ca2+-ATPase isoform effects on Ca2+ transients of embryonic chicken and neonatal rat cardiac myocytes. J. Physiol. 528, 53–63 (2000).
Zhai, J. et al. Cardiac-specific overexpression of a superinhibitory pentameric phospholamban mutant enhances inhibition of cardiac function in vivo. J. Biol. Chem. 275, 10538–10544 (2000).
Simmerman, H. K. & Jones, L. R. Phospholamban: protein structure, mechanism of action, and role in cardiac function. Physiol. Rev. 78, 921–947 (1998).
Liao, Z., Lockhead, D., Larson, E. D. & Proenza, C. Phosphorylation and modulation of hyperpolarization-activated HCN4 channels by protein kinase A in the mouse sinoatrial node. J. Gen. Physiol. 136, 247–258 (2010).
Mangoni, M. E. et al. Voltage-dependent calcium channels and cardiac pacemaker activity: from ionic currents to genes. Prog. Biophys. Mol. Biol. 90, 38–63 (2006).
McCormack, J. G., Halestrap, A. P. & Denton, R. M. Role of calcium ions in regulation of mammalian intramitochondrial metabolism. Physiol. Rev. 70, 391–425 (1990).
Rizzuto, R., De Stefani, D., Raffaello, A. & Mammucari, C. Mitochondria as sensors and regulators of calcium signalling. Nat. Rev. Mol. Cell. Biol. 13, 566–578 (2012).
Nagai, T., Sawano, A., Park, E. S. & Miyawaki, A. Circularly permuted green fluorescent proteins engineered to sense Ca2+. Proc. Natl Acad. Sci. USA 98, 3197–3202 (2001).
Territo, P. R., Mootha, V. K., French, S. A. & Balaban, R. S. Ca(2+) activation of heart mitochondrial oxidative phosphorylation: role of the F(0)/F(1)-ATPase. Am. J. Physiol. Cell Physiol. 278, C423–C435 (2000).
Wu, Y. et al. Calmodulin kinase II is required for fight or flight sinoatrial node physiology. Proc. Natl Acad. Sci. USA 106, 5972–5977 (2009).
Vinogradova, T. M., Bogdanov, K. Y. & Lakatta, E. G. beta-Adrenergic stimulation modulates ryanodine receptor Ca(2+) release during diastolic depolarization to accelerate pacemaker activity in rabbit sinoatrial nodal cells. Circ. Res. 90, 73–79 (2002).
Landolfi, B., Curci, S., Debellis, L., Pozzan, T. & Hofer, A. M. Ca2+ homeostasis in the agonist-sensitive internal store: functional interactions between mitochondria and the ER measured In situ in intact cells. J. Cell Biol. 142, 1235–1243 (1998).
Hofer, A. M., Schlue, W. R., Curci, S. & Machen, T. E. Spatial distribution and quantitation of free luminal [Ca] within the InsP3-sensitive internal store of individual BHK-21 cells: ion dependence of InsP3-induced Ca release and reloading. FASEB J. 9, 788–798 (1995).
Pan, X. et al. The physiological role of mitochondrial calcium revealed by mice lacking the mitochondrial calcium uniporter. Nat. Cell Biol. 15, 1464–1472 (2013).
Mangoni, M. E. & Nargeot, J. Properties of the hyperpolarization-activated current (I(f)) in isolated mouse sino-atrial cells. Cardiovasc. Res. 52, 51–64 (2001).
Rae, J., Cooper, K., Gates, P. & Watsky, M. Low access resistance perforated patch recordings using amphotericin B. J. Neurosci. Methods 37, 15–26 (1991).
DiFrancesco, D., Ferroni, A., Mazzanti, M. & Tromba, C. Properties of the hyperpolarizing-activated current (if) in cells isolated from the rabbit sino-atrial node. J. Physiol. 377, 61–88 (1986).
Cho, H. S., Takano, M. & Noma, A. The electrophysiological properties of spontaneously beating pacemaker cells isolated from mouse sinoatrial node. J. Physiol. 550, 169–180 (2003).
Baruscotti, M. et al. Deep bradycardia and heart block caused by inducible cardiac-specific knockout of the pacemaker channel gene Hcn4. Proc. Natl Acad. Sci. USA 108, 1705–1710 (2011).
Palmer, A. E. et al. Ca2+ indicators based on computationally redesigned calmodulin-peptide pairs. Chem. Biol. 13, 521–530 (2006).
Aon, M. A., Cortassa, S., Marban, E. & O'Rourke, B. Synchronized whole cell oscillations in mitochondrial metabolism triggered by a local release of reactive oxygen species in cardiac myocytes. J. Biol. Chem. 278, 44735–44744 (2003).
Eng, J., Lynch, R. M. & Balaban, R. S. Nicotinamide adenine dinucleotide fluorescence spectroscopy and imaging of isolated cardiac myocytes. Biophys. J. 55, 621–630 (1989).
Jiang, D., Zhao, L. & Clapham, D. E. Genome-wide RNAi screen identifies Letm1 as a mitochondrial Ca2+/H+ antiporter. Science 326, 144–147 (2009).
Feng, S. et al. Canonical transient receptor potential 3 channels regulate mitochondrial calcium uptake. Proc. Natl Acad. Sci. USA 110, 11011–11016 (2013).
Palty, R. et al. NCLX is an essential component of mitochondrial Na+/Ca2+ exchange. Proc. Natl Acad. Sci. USA 107, 436–441 (2010).
Kikuchi, K., McDonald, A. D., Sasano, T. & Donahue, J. K. Targeted modification of atrial electrophysiology by homogeneous transmural atrial gene transfer. Circulation 111, 264–270 (2005).
Joiner, M. L. et al. CaMKII determines mitochondrial stress responses in heart. Nature 491, 269–273 (2012).
Murphy, A.N., Bredesen, D.E., Cortopassi, G., Wang, E. & Fiskum, G. Bcl-2 potentiates the maximal calcium uptake capacity of neural cell mitochondria. Proc. Natl. Acad. Sci. USA 93, 9893–9898 (1996).
Purohit, A. et al. Oxidized Ca(2+)/calmodulin-dependent protein kinase II triggers atrial fibrillation. Circulation 128, 1748–1757 (2013).
Chen, B., Wu, Y., Mohler, P. J., Anderson, M. E. & Song, L. S. Local control of Ca2+-induced Ca2+ release in mouse sinoatrial node cells. J. Mol. Cell. Cardiol. 47, 706–715 (2009).
Weiss, R. M., Ohashi, M., Miller, J. D., Young, S. G. & Heistad, D. D. Calcific aortic valve stenosis in old hypercholesterolemic mice. Circulation 114, 2065–2069 (2006).
The authors thank Shawn Roach and Catherine E. Kiefe for graphic contributions, Rajan Sah, Peter M. Snyder and Leonid V. Zingman for their criticisms and suggestions, William J. Kutschke and Jinying Yang for technical support, Nicholas R. Wilson and Katrina Dion for experimental assistance and The University of Iowa Gene Transfer Vector Core for providing adenoviral agents. This work was supported by National Institutes of Health (NIH) Grants R01-HL079031, R01-HL096652, R01-HL070250 and R01-HL113001 to M.E.A., R01-HL089598 and R01-HL117641 to X.H.T.W. and F30-HL114258 to T.P.R.
Y.W. and M.E.A. are named inventors on a patent application claiming to control excessive heart rates by MCU inhibition.
Rights and permissions
About this article
Cite this article
Wu, Y., Rasmussen, T., Koval, O. et al. The mitochondrial uniporter controls fight or flight heart rate increases. Nat Commun 6, 6081 (2015). https://doi.org/10.1038/ncomms7081
This article is cited by
Mitochondrial Ca2+ uptake by the MCU facilitates pyramidal neuron excitability and metabolism during action potential firing
Communications Biology (2022)
Voltage-energized calcium-sensitive ATP production by mitochondria
Nature Metabolism (2019)
Tamoxifen-induced knockdown of the mitochondrial calcium uniporter in Thy1-expressing neurons protects mice from hypoxic/ischemic brain injury
Cell Death & Disease (2018)
The machineries, regulation and cellular functions of mitochondrial calcium
Nature Reviews Molecular Cell Biology (2018)
Mitochondrial calcium uptake in organ physiology: from molecular mechanism to animal models
Pflügers Archiv - European Journal of Physiology (2018)
By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.