Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Letter
  • Published:

Voltage-energized calcium-sensitive ATP production by mitochondria

Nature Metabolism volume 1, pages 975–984 (2019)Cite this article

Subjects

An Author Correction to this article was published on 30 October 2019

This article has been updated

Abstract

The regulation of ATP production by mitochondria, crucial for multicellular life, is poorly understood. Here, we investigate the molecular controls of this process in the heart and provide a framework for its Ca2+-dependent regulation. We find that the entry of Ca2+ into the matrix through the mitochondrial calcium uniporter (MCU) in the heart has neither an apparent cytosolic Ca2+ threshold nor a gating function, and guides ATP production by its influence on the inner mitochondrial membrane (IMM) potential, ΔΨm. This regulation occurs through matrix Ca2+-dependent modulation of pyruvate and glutamate dehydrogenase activity and not through any effect of Ca2+ on ATP synthase or on electron transport chain complexes II, III or IV. Examining the ΔΨm dependence of ATP production over the range of −60 mV to −170 mV in detail reveals that cardiac ATP synthase has a voltage dependence that distinguishes it fundamentally from the previous standard, the bacterial ATP synthase. Cardiac ATP synthase operates with a different ΔΨm threshold for ATP production than bacterial ATP synthase and reveals a concave-upward shape without saturation. Skeletal muscle MCU Ca2+ flux, while also having no apparent cytosolic Ca2+ threshold, is substantially different from the cardiac MCU, yet the ATP synthase voltage dependence in skeletal muscle is identical to that in the heart. These results suggest that, while the conduction of cytosolic Ca2+ signals through the MCU appears to be tissue dependent, as shown by earlier work1, the control of ATP synthase by ΔΨm appears to be broadly consistent among tissues but is clearly different from that in bacteria.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on SpringerLink
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: [Ca2+]m-sensitive ATP production by mitochondria.
Fig. 2: [Ca2+]m control of ΔΨm.
Fig. 3: [Ca2+]m dynamics and MCU Ca2+ conductance.
Fig. 4: ΔΨm control of ATP production.

Similar content being viewed by others

Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Change history

  • 30 October 2019

    An amendment to this paper has been published and can be accessed via a link at the top of the paper.

References

  1. Fieni, F., Lee, S. B., Jan, Y. N. & Kirichok, Y. Activity of the mitochondrial calcium uniporter varies greatly between tissues. Nat. Commun. 3, 1317 (2012).

    Article  Google Scholar 

  2. Jacobus, W. E. Respiratory control and the integration of heart high-energy phosphate metabolism by mitochondrial creatine kinase. Annu. Rev. Physiol. 47, 707–725 (1985).

    Article  CAS  Google Scholar 

  3. Wang, Z. et al. Specific metabolic rates of major organs and tissues across adulthood: evaluation by mechanistic model of resting energy expenditure. Am. J. Clin. Nutr. 92, 1369–1377 (2010).

    Article  CAS  Google Scholar 

  4. Murphy, E. et al. Mitochondrial function, biology, and role in disease: a scientific statement from the American Heart Association. Circ. Res. 118, 1960–1991 (2016).

    Article  CAS  Google Scholar 

  5. DiMauro, S. & Schon, E. A. Mitochondrial respiratory-chain diseases. N. Engl. J. Med. 348, 2656–2668 (2003).

    Article  CAS  Google Scholar 

  6. Latorre-Pellicer, A. et al. Mitochondrial and nuclear DNA matching shapes metabolism and healthy ageing. Nature 535, 561–565 (2016).

    Article  CAS  Google Scholar 

  7. Williams, G. S., Boyman, L., Chikando, A. C., Khairallah, R. J. & Lederer, W. J. Mitochondrial calcium uptake. Proc. Natl Acad. Sci. USA 110, 10479–10486 (2013).

    Article  CAS  Google Scholar 

  8. Boyman, L., Williams, G. S. & Lederer, W. J. The growing importance of mitochondrial calcium in health and disease. Proc. Natl Acad. Sci. USA 112, 11150–11151 (2015).

    Article  CAS  Google Scholar 

  9. Chance, B. & Williams, G. R. Respiratory enzymes in oxidative phosphorylation. I. Kinetics of oxygen utilization. J. Biol. Chem. 217, 383–393 (1955).

    CAS  PubMed  Google Scholar 

  10. Klingenberg, M. The ADP-ATP translocation in mitochondria, a membrane potential controlled transport. J. Membr. Biol. 56, 97–105 (1980).

    Article  CAS  Google Scholar 

  11. Denton, R. M., Randle, P. J. & Martin, B. R. Stimulation by calcium ions of pyruvate dehydrogenase phosphate phosphatase. Biochem. J. 128, 161–163 (1972).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. McCormack, J. G. & Denton, R. M. The effects of calcium ions and adenine nucleotides on the activity of pig heart 2-oxoglutarate dehydrogenase complex. Biochem. J. 180, 533–544 (1979).

    Article  CAS  Google Scholar 

  13. Glancy, B. & Balaban, R. S. Role of mitochondrial Ca2+ in the regulation of cellular energetics. Biochemistry 51, 2959–2973 (2012).

    Article  CAS  Google Scholar 

  14. McCormack, J. G. & Denton, R. M. Mitochondrial Ca2+ transport and the role of intramitochondrial Ca2+ in the regulation of energy metabolism. Dev. Neurosci. 15, 165–173 (1993).

    Article  CAS  Google Scholar 

  15. Jian, Z. et al. Mechanochemotransduction during cardiomyocyte contraction is mediated by localized nitric oxide signaling. Sci. Signal 7, ra27 (2014).

    Article  Google Scholar 

  16. Mitchell, P. Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191, 144–148 (1961).

    Article  CAS  Google Scholar 

  17. Brandes, R., Maier, L. S. & Bers, D. M. Regulation of mitochondrial [NADH] by cytosolic [Ca2+] and work in trabeculae from hypertrophic and normal rat hearts. Circ. Res. 82, 1189–1198 (1998).

    Article  CAS  Google Scholar 

  18. Kirichok, Y., Krapivinsky, G. & Clapham, D. E. The mitochondrial calcium uniporter is a highly selective ion channel. Nature 427, 360–364 (2004).

    Article  CAS  Google Scholar 

  19. Bick, A. G., Calvo, S. E. & Mootha, V. K. Evolutionary diversity of the mitochondrial calcium uniporter. Science 336, 886 (2012).

    Article  CAS  Google Scholar 

  20. Luongo, T. S. et al. The mitochondrial calcium uniporter matches energetic supply with cardiac workload during stress and modulates permeability transition. Cell Rep. 12, 23–34 (2015).

    Article  CAS  Google Scholar 

  21. Kwong, J. Q. et al. The mitochondrial calcium uniporter selectively matches metabolic output to acute contractile stress in the heart. Cell Rep. 12, 15–22 (2015).

    Article  CAS  Google Scholar 

  22. Wu, Y. et al. The mitochondrial uniporter controls fight or flight heart rate increases. Nat. Commun. 6, 6081 (2015).

    Article  CAS  Google Scholar 

  23. Mallilankaraman, K. et al. MICU1 is an essential gatekeeper for MCU-mediated mitochondrial Ca(2+) uptake that regulates cell survival. Cell 151, 630–644 (2012).

    Article  CAS  Google Scholar 

  24. Antony, A. N. et al. MICU1 regulation of mitochondrial Ca(2+) uptake dictates survival and tissue regeneration. Nat. Commun 7, 10955 (2016).

    Article  CAS  Google Scholar 

  25. Kaim, G. & Dimroth, P. ATP synthesis by F-type ATP synthase is obligatorily dependent on the transmembrane voltage. EMBO J. 18, 4118–4127 (1999).

    Article  CAS  Google Scholar 

  26. Nguyen, M. H. & Jafri, M. S. Mitochondrial calcium signaling and energy metabolism. Ann. N Y Acad. Sci. 1047, 127–137 (2005).

    Article  CAS  Google Scholar 

  27. Dimroth, P., von Ballmoos, C., Meier, T. & Kaim, G. Electrical power fuels rotary ATP synthase. Structure 11, 1469–1473 (2003).

    Article  CAS  Google Scholar 

  28. Anandakrishnan, R., Zhang, Z., Donovan-Maiye, R. & Zuckerman, D. M. Biophysical comparison of ATP synthesis mechanisms shows a kinetic advantage for the rotary process. Proc. Natl Acad. Sci. USA 113, 11220–11225 (2016).

    Article  CAS  Google Scholar 

  29. Clapham, D. E. Calcium signaling. Cell 131, 1047–1058 (2007).

    Article  CAS  Google Scholar 

  30. Cheng, H. & Lederer, W. J. Calcium sparks. Physiol. Rev. 88, 1491–1545 (2008).

    Article  CAS  Google Scholar 

  31. Eberhard, M. & Erne, P. Analysis of calcium binding to alpha-lactalbumin using a fluorescent calcium indicator. Eur. J. Biochem. 202, 1333–1338 (1991).

    Article  CAS  Google Scholar 

  32. Scaduto, R. C. Jr. & Grotyohann, L. W. Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives. Biophys. J. 76, 469–477 (1999).

    Article  CAS  Google Scholar 

  33. 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).

    Article  Google Scholar 

  34. Burks, S. R. et al. (2)H,(15)N-substituted nitroxides as sensitive probes for electron paramagnetic resonance imaging. J. Org. Chem. 75, 6463–6467 (2010).

    Article  CAS  Google Scholar 

  35. Rosen, G. M., Burks, S. R., Kohr, M. J. & Kao, J. P. Synthesis and biological testing of aminoxyls designed for long-term retention by living cells. Org. Biomol. Chem. 3, 645–648 (2005).

    Article  CAS  Google Scholar 

  36. Terhzaz, S. et al. Differential gel electrophoresis and transgenic mitochondrial calcium reporters demonstrate spatiotemporal filtering in calcium control of mitochondria. J. Biol. Chem. 281, 18849–18858 (2006).

    Article  CAS  Google Scholar 

  37. Boyman, L. et al. Calcium movement in cardiac mitochondria. Biophys. J. 107, 1289–1301 (2014).

    Article  CAS  Google Scholar 

Download references

Acknowledgements

We thank B.M. Polster, C.W. Ward, G.S.B. Williams, R.J . Khairallah and M. Karbowski for helpful suggestions. This research was supported by American Heart Association grants SDG 15SDG22100002 (to L.B.) and 16PRE31030023 (to A.P.W.); by NIH R01 HL106056, R01 HL105239, U01 HL116321, 1R01HL142290, 1 R01 HL140934 and 1R01 AR071618 (to W.J.L.); by the Medical Scientist Training Program and Training Program in Integrative Membrane Biology, NIH 2T32GM092237-06 and 5T32GM008181-28 (to A.P.W.).

Author information

Authors and Affiliations

  1. Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD, USA

    Andrew P. Wescott, Joseph P. Y. Kao, W. Jonathan Lederer & Liron Boyman

  2. Department of Physiology, University of Maryland School of Medicine, Baltimore, MD, USA

    Andrew P. Wescott, Joseph P. Y. Kao, W. Jonathan Lederer & Liron Boyman

Authors
  1. Andrew P. Wescott
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Joseph P. Y. Kao
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. W. Jonathan Lederer
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Liron Boyman
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

A.P.W. and L.B. were involved with all experiments, data collection and data analysis; A.P.W., L.B. and W.J.L. contributed to study design, data interpretation and writing the paper; J.P.Y.K. synthesized the four-armed PEG-boronic acid and the diAM-succinate. All authors edited the manuscript.

Corresponding author

Correspondence to Liron Boyman.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data

Extended Data Fig. 1 Mitochondrial ATP production, NADH, voltage and pH.

a, Timecourse of firefly luciferase luminescence signal measured from nine wells after addition of ATP to each well. In this calibration procedure, different amounts of [ATP] are added to each of the nine wells as indicated to the right of each line ([ATP]added in μM). The inset shows the measured luminescence versus [ATP]added. Purple shaded area highlights the ‘Working Range’ in which the luminescence signal is a linear function of [ATP]. b, Measurements of [ATP] produced by isolated mitochondria. The calibration procedure shown in a is used to convert the measured luminescence signal to [ATP]. The inset shows the measured mitochondrial matrix free Ca2+ concentration ([Ca2+]m) associated with the ATP measurements in b. c, ATP production rate based on the linear fit to measurements in b and scaled to units of μmol per litre cytosol per second (μM s−1, scaling is based on 80 g mitochondrial protein per litre cardiomyocyte cytosol, for more details see main methods section and Williams et. al.7) . d, The increase in [NADH] at high [Ca2+]m (>2 μM) relative to [NADH] at low [Ca2+]m (<200 nM) using the indicated combination of carbon substrates (P&M, pyruvate and malate; G&M, glutamate and malate; PC&M, palmitoylcarnitine and malate, n = 4 isolated mitochondria preparations per group, *P < 0.05 by two-sided t-test). Data are mean ± s.e.m. [Ca2+]m was measured with Rhod-2. [NADH] measurements were carried out with a luminescence assay kit (Promega, USA, for additional details see Supplementary Methods 1.4). e, Measured TMRM fluorescence ratio (F573/F46) over the maximal fluorescence ratio from the dataset. Mitochondria are exposed to 2,4-dinitrophenol ([DNP] in μM) as indicated. f, Measured extra-mitochondrial TMRM concentration. g, The mitochondrial inner membrane potential (ΔΨm) in mV is obtained from the measurements in f according to the method by Scaduto & Grotyohann32. h, ΔΨm and its corresponding TMRM fluorescence ratio. Linear fit lines are as indicated in the inset. The calibration results shown in panels e–h were verified repeatedly on a daily basis with similar results obtained. i, Excitation and emission spectra of mitochondria loaded with BCECF (2′,7′-bis(2-carboxyethyl)-5(6)-carboxyfluorescein acetoxymethyl ester). 15 independents measurements are shown at the indicated pH levels (1 μM FCCP is used to equilibrate pHm and the extra-mitochondrial buffer pH) with the similar spectrum shown. j, Measured fluorescence ratio from BCECF-loaded mitochondria at the indicated mitochondrial pH values (pHm). The calibration was verified in two mitochondrial preps, with similar results. k, The pHm measurements following exposure to sodium acetate at the indicated concentrations. Data are mean ± s.e.m. (results are from 3 independent experiments in each of the indicated 7 concentrations of sodium acetate).

Extended Data Fig. 2 Calibration of fluorescence measurements with Ca2+ indicators.

a, Fluo-4 Ca2+ titration curve. Shown is the fraction of Ca2+-bound fluo-4 bound at the indicated added [Ca2+]. Each point is a triplicate average. Titration curves are carried out in the indicated buffers. b, Fluo-4FF Ca2+ titration curve. Shown is the fraction of Ca2+-bound fluo-4FF at the indicated added [Ca2+]. Each point is a triplicate average. c, Rhod-2 Ca2+ titration curve. Shown is the fraction of Ca2+-bound rhod-2 at the indicated added [Ca2+]. Each point is a triplicate average. Titration is done in the absence and presence of PVP (polyvinylpyrrolidone) in the assay buffer. d, Ca2+ titration curve for fura-2AM- or rhod-2AM-loaded mitochondria. Shown is the fraction of Ca2+-bound fura-2 or rhod-2 at the indicated free Ca2+ concentration in the mitochondrial matrix ([Ca2+]m). Each point is a triplicate average. 1 μM FCCP and 1 μM of the Ca2+ ionophore ionomycin are used to equilibrate [Ca2+]m with the extra-mitochondrial free [Ca2+] (that is, [Ca2+]extra,free). The [Ca2+]extra,free is measured with fluo-5N. Equations fit to the data for a–d are detailed in the main methods section.

Extended Data Fig. 3 Measurements of mitochondrial ATP production when the MCU is blocked by RU360.

a, Measurements of [Ca2+]m in isolated cardiac mitochondria plotted as a function of the measured free extramitochondrial [Ca2+] (that is, [Ca2+]extra,free) in the presence of the selective MCU blocker RU360 (5 μM). Grey circles are measurements taken in the presence of pyruvate (1 mM) and malate (0.5 mM). n = 12 independent experiments. Orange circles are measurements done in the presence of glutamate (1 mM) and malate (0.5 mM). n = 12 independent experiments. [Ca2+]m was measured with fura-2 loaded into the mitochondrial matrix via its acetoxymethyl (AM) ester, [Ca2+]extra,free was measured with fluo-4 in the extra mitochondrial buffer. b, Measurements of ATP production plotted as a function of the measured [Ca2+]extra,free. The measurements of ATP production rates are normalized to the measurements at nominally zero [Ca2+]extra,free. c, ATP production at the indicated measured [Ca2+]extra,free (from experiments in a and b). Grey bars (left) are ATP production with pyruvate and malate. Orange bars (right) are ATP production with glutamate and malate. Data are mean ± s.e.m. n = 3 isolated mitochondria preparations per group (*P < 0.05 by one-way two tailed ANOVA with Bonferroni correction).

Extended Data Fig. 4 Synthesis of 4-armed PEG-boronic acid.

Abbreviations used: HBTU, 1-[bis(dimethylamino)methylene]-1H-benzotriazolium hexafluorophosphate 3-oxide; DIPEA, diisopropylethylamine; DMF, N,N-dimethylformamide. The detailed description of the synthesis procedure is in the methods section.

Extended Data Fig. 5 ΔΨm kinetics during mitochondrial ATP production.

a, Time-dependent stopped-flow measurement of ΔΨm (upper panel) and of the corresponding [Ca2+]m (lower panel). In this protocol (no. 1), mitochondria were incubated with increasing extra-mitochondrial free Ca2+ and at t = 0, 500 μM [ADP] was added to the mitochondrial mix. Time-dependent depolarization of ΔΨm is shown as is the near steady-state of [Ca2+]m. Black line: [Ca2+]m= 50 nM (n = 8); turquoise: [Ca2+]m= 480 nM (n = 3); light blue: [Ca2+]m= 750 nM (n = 6); grey-blue: [Ca2+]m= 1.2 μM (n = 8); navy blue: [Ca2+]m= 1.7 μM (n = 4). b, Same as a, but in this protocol (no. 2), [Ca2+]m and [ADP] (500 μM) were increased simultaneously at t = 0 (salmon-coloured line, n = 7). The injected Ca2+ was set so that the [Ca2+]m achieved a level between 1.5 and 2 μM at 20 s. c, Same as b but in this protocol (#3), [ADP] (500 μM) rises 10 s before [Ca2+]m was increased at t = 0 (grey line, n = 6). The injected Ca2+ was set so that the [Ca2+]m achieved a level between 1.5 and 2 μM at 20 s, grey line. In a–c, the sample size (n) represents the number of independent experiments. d, The magnitude of ΔΨm depolarization in experiments a–c. The sample size is the same as in panels a–c. (* and # denotes statistical comparisons to black and beige data, respectively). e, Exponential rate constant of ΔΨm depolarization in experiments a and b. The sample size is the same as in panels a and b. In d and e, data are mean ± s.e.m.*P < 0.05, **P < 0.01, ***P < 0.001 by one-way two-tailed ANOVA with Bonferroni correction.

Extended Data Fig. 6 Stopped-flow measurements of MCU Ca2+ flux and its driving force.

a,e, Representative stopped-flow time-dependent measurements of extra-mitochondrial free Ca2+ (that is, [Ca2+]extra,free). Mitochondria in uptake assay buffer (uAB) with low [Ca2+]extra,free (<100 nM) are mixed with uAB buffer with added [Ca2+] 1 ms before fluorescence measurements begin. The levels of added [Ca2+] are set so that at the beginning of the measurements the [Ca2+] added will be as indicated in the inset. In a, [Ca2+]extra,free is measured with fluo-4 in the uAB and in e with Fluo-4FF. b,f, The corresponding time-dependent measurements of matrix free Ca2+ (that is, [Ca2+]m). Insets showing the corresponding time-dependent measurements of ΔΨm. c,g, The corresponding time-dependent measurements of total extra-mitochondrial Ca2+ ([Ca2+]extra,Total). The [Ca2+]extra,Total is obtained from the fluo-4 or fluo-4FF signals (for more details see the Methods). d,h, MCU Ca2+ influx (Jmcu) is the first derivative of the [Ca2+]extra,Total. The Jmcu is scaled to units of μmol per litre cytosol per second (μM s−1, scaling is based on 80 g mitochondrial protein per litre cardiomyocyte cytosol, for more details see main methods section and Williams et. al.7). The shown stopped flow experiments were repeated independently 63 times with similar results at each [Ca2+]extra,free as indicated in Fig. 3a.

Extended Data Fig. 7 Characterization of MityCam, a mitochondrially targeted Ca2+-sensitive fluorescent-protein probe expressed in heart muscle cells.

a, MityCam fluorescence versus [Ca2+]m. Note that Ca2+ binding decreases MityCam fluorescence. Measurements are in saponin-permeabilized cardiomyocytes; [Ca2+]m is set using the Ca2+ ionophore ionomycin (2 μM). The extracellular (bath) solution contains Rhod-2 (tripotassium salt, cell-impermeant) to measure the bath [Ca2+], a proton ionophore carbonyl cyanide m-chlorophenyl hydrazone (CCCP, 500 nM) to set the mitochondrial pH, rotenone (400 nM) to block the ETC and the production of reactive oxygen species, and oligomycin (5 mM) to block reverse-mode consumption of ATP by the ATP synthase, pH 7.8. Fit curve is a single-site binding model (n = 30 cells). b, Top: Confocal line-image from an intact cardiomyocyte expressing MityCam. Bottom: The time-course of changes in [Ca2+]m from the confocal fluorescence measurements. Caffeine (10 mM) was applied for 10 s via a local micro-perfusion system to rapidly trigger Ca2+ release from the sarcoplasmic reticulum at the indicated times (highlighted with grey shading). The experiment was repeated independently with n = 10 cells with similar results. c, The average time-course of changes in [Ca2+]m following caffeine applications. d, Confocal line-image from an intact cardiomyocyte expressing MityCam in a bath solution devoid of Ca2+ (chelated by 1 mM EGTA) and treated with thapsigargin (1 μM) for 10 min prior to imaging to deplete the sarcoplasmic reticulum of Ca2+. Top panel shows MityCam fluorescence; middle panel shows the fluorescence of sulforhodamine (sulforhodamine is included in the micro-perfusion solution to indicate the exact duration of 10 mM caffeine application). Lower panel shows the time-course of changes in [Ca2+]m from the confocal fluorescence measurements. Note that throughout the entire time course of the experiment the extracellular solution is devoid of Ca2+ and also contains 1 μM thapsigargin. The experiment was repeated independently with n = 11 cells with similar results. e, Confocal images of cardiac mitochondria isolated from MityCam expressing cardiomyocytes. f, Measurements of [Ca2+]m from isolated single mitochondria expressing MityCam. Top panel (i) shows MityCam fluorescence; lower panel (ii) shows sulforhodamine fluorescence, which indicates the duration of micro-perfusion of 100 μM Ca2+ (see bars above the top panel). Mitochondria isolated from MityCam-expressing cardiomyocytes are adhered to a glass coverslip for confocal microscopy measurements. Rapid step (2–3 ms rise time) of [Ca2+] from 0 (1 mM EGTA) to 100 μM is achieved via a micro-perfusion system. Sulforhodamine is included in the solution applied via the micro-perfusion system to indicate when a mitochondrion is exposed to the solution containing 100 μM Ca2+. The experiment in e and f were repeated independently with n = 18 mitochondria with similar results. g, The average time-course of changes in [Ca2+]m following the step increase of Ca2+ from 0 to 100 μM (black line). h, Same as g but for comparison to another Ca2+ sensitive fluorescent indicator, MityCam-expressing mitochondria are also loaded with the fluorescent Ca2+ indicator Rhod-2 via its acetoxymethyl (AM) ester form (Rhod-2 AM). To accelerate the rate of [Ca2+]m rise [Ca2+] is raised to 1 mM, all solutions also contain ionomycin (5 μM), FCCP (5 μM), oligomycin (1 μM), pH = 7.8. Green trace for MityCam (n = 12 mitochondria) and red for Rhod-2 (n = 9 mitochondria). The time at which 1 μM [Ca2+]m is measured is indicated by arrows, this time point is obtained by converting the shown fit lines to units of μM [Ca2+]m. For additional details see Boyman et al., 2014 (ref. 37).

Extended Data Fig. 8 MCU conductance and voltage dependence of ATP production in heart and skeletal muscle.

a, Measurement of the MCU-dependent Ca2+ influx (Jmcu) (nmole mg−1 s−1) in cardiac mitochondria (green circles, from Fig. 3), skeletal muscle (black circles), and skeletal muscle with Ru360 (5 μM) (red circles) is plotted as a function of measured [Ca2+]i (n = 63, n = 87, and n = 12 independent experiments, respectively, each with [Ca2+]i, [Ca2+]m and ΔΨm measured). Linear least-squares fit to the heart mitochondria data is shown (slope = 0.015). b, MCU conductance (G) for each measurement shown in a normalized to the minimal conductance (Gmin) of the cardiac dataset (G/Gmin) plotted as a function of [Ca2+]i. Linear least-squares fit line to the heart mitochondria data is shown (slope = 6.1). c, Relative number of open MCUs per mitochondrion plotted as a function of [Ca2+]I, taken from b after dividing by the [Ca2+]i-dependent unitary conductance of MCU and normalized to the minimal number of open MCUs of the cardiac dataset. Linear least-squares fit to the heart mitochondria data is shown (slope = 0.051, intercept = 3.3). For skeletal muscle data under [Ca2+]i of 1.5 μM the measurements were done using stopped flow as described in Extended Data Fig. 5. Jmcu at [Ca2+]i above 1.5 μM was collected using a multi-well-plate reader with ΔΨm set using a K+ gradient and the K+ ionophore valinomycin. Skeletal muscle data is fit to a modified Hill equation with a K0.5 of 7.9 μM and a Hill coefficient of 2.95. d, The dependence of ATP production on ΔΨm in the absence of carbon substrates and at [Ca2+]m < 200 nM. The measurements of ATP production rates are normalized to the minimal production rate of each dataset. Measurements from heart mitochondria are shown in green circles (n = 77, replotted from Fig. 4a), the measurements from skeletal muscle mitochondria are in shown in black circles (n = 45 independent experiments). ΔΨm was set by using a fixed K+ gradient and the K+ ionophore valinomycin (see Methods).

Supplementary information

Supplementary Information

Supplementary Figs. 1 and 2, and Discussion

Reporting Summary

Supplementary Table

Supplementary Tables containing statistical information

Supplementary Video 1 Video-based measurements of sarcomere length from cardiomyocyte contracting in PVA hydrogel

Video recording of contracting cardiomyocyte embedded in and cross-linked to the PVA hydrogel (captured at 300 Hz frame-rate). Dynamic video-based sarcomere length measurements in μm are carried out from the highlighted black rectangular frame with the 900B:VSL system (Aurora Scientific, Canada). Field electrical stimulation (40 Volt cm–1) to trigger contraction at either 1 Hz or 8 Hz was done using MyoPacer (Ion Optix, USA).

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Wescott, A.P., Kao, J.P.Y., Lederer, W.J. et al. Voltage-energized calcium-sensitive ATP production by mitochondria. Nat Metab 1, 975–984 (2019). https://doi.org/10.1038/s42255-019-0126-8

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s42255-019-0126-8

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing