Mitochondrial energetic adaptations encompass a plethora of conserved processes that maintain cell and organismal fitness and survival in the changing environment by adjusting the respiratory capacity of mitochondria. These mitochondrial responses are governed by general principles of regulatory biology exemplified by changes in gene expression, protein translation, protein complex formation, transmembrane transport, enzymatic activities and metabolite levels. These changes can promote mitochondrial biogenesis and membrane dynamics that in turn support mitochondrial respiration. The main regulatory components of mitochondrial energetic adaptation include: the transcription coactivator peroxisome proliferator-activated receptor-γ (PPARγ) coactivator 1α (PGC1α) and associated transcription factors; mTOR and endoplasmic reticulum stress signalling; TOM70-dependent mitochondrial protein import; the cristae remodelling factors, including mitochondrial contact site and cristae organizing system (MICOS) and OPA1; lipid remodelling; and the assembly and metabolite-dependent regulation of respiratory complexes. These adaptive molecular and structural mechanisms increase respiration to maintain basic processes specific to cell types and tissues. Failure to execute these regulatory responses causes cell damage and inflammation or senescence, compromising cell survival and the ability to adapt to energetically demanding conditions. Thus, mitochondrial adaptive cellular processes are important for physiological responses, including to nutrient availability, temperature and physical activity, and their failure leads to diseases associated with mitochondrial dysfunction such as metabolic and age-associated diseases and cancer.
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Gilkerson, R. W., Selker, J. M. L. & Capaldi, R. A. The cristal membrane of mitochondria is the principal site of oxidative phosphorylation. FEBS Lett. 546, 355–358 (2003).
Deshpande, O. A. & Mohiuddin, S. S. Biochemistry, Oxidative Phophorylation (StatPearls, 2020).
Walker, J. E. The ATP synthase: the understood, the uncertain and the unknown. Biochem. Soc. Trans. 41, 1–16 (2013).
Enerbäck, S. et al. Mice lacking mitochondrial uncoupling protein are cold-sensitive but not obese. Nature 387, 90–94 (1997).
Kazak, L. et al. A creatine-driven substrate cycle enhances energy expenditure and thermogenesis in beige fat. Cell 163, 643–655 (2015).
Nedergaard, J. & Cannon, B. Brown adipose tissue as a heat-producing thermoeffector. Handb. Clin. Neurol. 156, 137–152 (2018).
Lowell, B. B. & Spiegelman, B. M. Towards a molecular understanding of adaptive thermogenesis. Nature 404, 652–660 (2000).
Chouchani, E. T. et al. Ischaemic accumulation of succinate controls reperfusion injury through mitochondrial ROS. Nature 515, 431–435 (2014).
Granger, D. N. & Kvietys, P. R. Reperfusion injury and reactive oxygen species: the evolution of a concept. Redox Biol. 6, 524–551 (2015).
Chouchani, E. T. et al. Mitochondrial ROS regulate thermogenic energy expenditure and sulfenylation of UCP1. Nature 532, 112–116 (2016).
Bock, F. J. & Tait, S. W. G. Mitochondria as multifaceted regulators of cell death. Nat. Rev. Mol. Cell Biol. 21, 85–100 (2020).
Giorgi, C., Marchi, S. & Pinton, P. The machineries, regulation and cellular functions of mitochondrial calcium. Nat. Rev. Mol. Cell Biol. 19, 713–730 (2018).
Spinelli, J. B. & Haigis, M. C. The multifaceted contributions of mitochondria to cellular metabolism. Nat. Cell Biol. 20, 745–754 (2018).
Stephan, T. et al. MICOS assembly controls mitochondrial inner membrane remodeling and crista junction redistribution to mediate cristae formation. EMBO J. 39, e104105 (2020). This article, along with Kondadi et al. (2020), dissects the involvement of several MICOS subunits in cristae remodelling and dynamics using advanced microscopy techniques.
Frezza, C. et al. OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion. Cell 126, 177–189 (2006).
Civiletto, G. et al. Opa1 overexpression ameliorates the phenotype of two mitochondrial disease mouse models. Cell Metab. 21, 845–854 (2015).
Sustarsic, E. G. et al. Cardiolipin synthesis in brown and beige fat mitochondria is essential for systemic energy homeostasis. Cell Metab. 28, 159–174.e11 (2018). This work uncovers the lipidomic alterations in beige and brown adipose tissues essential for thermogenesis in mice.
Bennett, C. F. et al. Peroxisomal-derived ether phospholipids link nucleotides to respirasome assembly. Nat. Chem. Biol. 17, 703–710 (2021).
Kondadi, A. K. et al. Cristae undergo continuous cycles of membrane remodelling in a MICOS -dependent manner. EMBO Rep. 21, e49776 (2020).
Balsa, E. et al. ER and nutrient stress promote assembly of respiratory chain supercomplexes through the PERK-eIF2α axis. Mol. Cell 74, 877–890.e6 (2019).
Latorre-Muro, P. et al. A cold-stress-inducible PERK/OGT axis controls TOM70-assisted mitochondrial protein import and cristae formation. Cell Metab. 33, 598–614.e7 (2021). This article describes regulation of cristae formation by ER stress responses, along with Kato et al. (2020).
Giacomello, M., Pyakurel, A., Glytsou, C. & Scorrano, L. The cell biology of mitochondrial membrane dynamics. Nat. Rev. Mol. Cell Biol. 21, 204–224 (2020).
Puigserver, P. et al. A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell 92, 829–839 (1998). This is the initial report detailing PGC1α as a transcriptional coactivator that binds nuclear receptors and promotes expression of mitochondrial genes.
Wu, Z. et al. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell 98, 115–124 (1999).
Holloszy, J. O. Biochemical adaptations in muscle. Effects of exercise on mitochondrial oxygen uptake and respiratory enzyme activity in skeletal muscle. J. Biol. Chem. 242, 2278–2282 (1967).
Handschin, C. et al. Skeletal muscle fiber-type switching, exercise intolerance, and myopathy in PGC-1α muscle-specific knock-out animals. J. Biol. Chem. 282, 30014–30021 (2007).
Couvillion, M. T., Soto, I. C., Shipkovenska, G. & Churchman, L. S. Synchronized mitochondrial and cytosolic translation programs. Nature 533, 499–503 (2016).
Soto, I. et al. Balanced mitochondrial and cytosolic translatomes underlie the biogenesis of human respiratory complexes. Preprint at bioRxiv https://doi.org/10.1101/2021.05.31.446345 (2021).
Wang, C. et al. MITRAC15/COA1 promotes mitochondrial translation in a ND2 ribosome–nascent chain complex. EMBO Rep 21, e48833 (2020).
Richter-Dennerlein, R. et al. Mitochondrial protein synthesis adapts to influx of nuclear-encoded protein. Cell 167, 471–483.e10 (2016). This article describes plasticity in translational rate from membrane-associated human mitochondrial ribosome–mRNA complexes depending on the import of nuclear-encoded complex IV subunits.
Mick, D. U. et al. MITRAC links mitochondrial protein translocation to respiratory-chain assembly and translational regulation. Cell 151, 1528–1541 (2012).
Rath, S. et al. MitoCarta3.0: an updated mitochondrial proteome now with sub-organelle localization and pathway annotations. Nucleic Acids Res. 49, D1541–D1547 (2021).
Rensvold, J. W. et al. Complementary RNA and protein profiling identifies iron as a key regulator of mitochondrial biogenesis. Cell Rep. 3, 237–245 (2013).
Holloszy, J. O. Regulation by exercise of skeletal muscle content of mitochondria and GLUT4. J. Physiol. Pharmacol. 59, 5–18 (2008).
Lin, J. et al. Transcriptional co-activator PGC-1α drives the formation of slow-twitch muscle fibres. Nature 418, 797–801 (2002).
Lai, L. et al. Transcriptional coactivators PGC-lα and PGC-lβ control overlapping programs required for perinatal maturation of the heart. Genes Dev. 22, 1948–1961 (2008).
Ciron, C. et al. PGC-1α activity in nigral dopamine neurons determines vulnerability to α-synuclein. Acta Neuropathol. Commun. 3, 16 (2015).
Jiang, H. et al. Adult conditional knockout of PGC-1α leads to loss of dopamine neurons. eNeuro 3, ENEURO.0183-16.2016 (2016).
Tran, M. T. et al. PGC1α drives NAD biosynthesis linking oxidative metabolism to renal protection. Nature 531, 528–532 (2016).
Mutlu, B. & Puigserver, P. GCN5 acetyltransferase in cellular energetic and metabolic processes. Biochim. Biophys. Acta Gene Regul. Mech. 1864, 194626 (2021).
Luo, C., Widlund, H. R. & Puigserver, P. PGC-1 coactivators: shepherding the mitochondrial biogenesis of tumors. Trends Cancer 2, 619–631 (2016).
Dominy, J. E. & Puigserver, P. Mitochondrial biogenesis through activation of nuclear signaling proteins. Cold Spring Harb. Perspect. Biol. https://doi.org/10.1101/cshperspect.a015008 (2013).
Virbasius, J. V. & Scarpulla, R. C. Activation of the human mitochondrial transcription factor A gene by nuclear respiratory factors: A potential regulatory link between nuclear and mitochondrial gene expression in organelle biogenesis. Proc. Natl Acad. Sci. USA 91, 1309–1313 (1994).
Brown, E. L. et al. PGC-1α and PGC-1β increase protein synthesis via ERRα in C2C12 myotubes. Front. Physiol. 9, 1336 (2018).
Handschin, C. et al. Nutritional regulation of hepatic heme biosynthesis and porphyria through PGC-1α. Cell 122, 505–515 (2005).
Yoon, J. C. et al. Control of hepatic gluconeogenesis through the transcriptional coactivator PGC-1. Nature 413, 131–138 (2001).
Blättler, S. M. et al. Defective mitochondrial morphology and bioenergetic function in mice lacking the transcription factor Yin Yang 1 in skeletal muscle. Mol. Cell. Biol. 32, 3333–3346 (2012).
Scarpulla, R. C., Vega, R. B. & Kelly, D. P. Transcriptional integration of mitochondrial biogenesis. Trends Endocrinol. Metab. 23, 459–466 (2012).
Tavares, C. D. J. et al. Transcriptome-wide analysis of PGC-1α-binding RNAs identifies genes linked to glucagon metabolic action. Proc. Natl Acad. Sci. USA 117, 22204–22213 (2020).
Schreiber, S. N. et al. The estrogen-related receptor α (ERRα) functions in PPARγ coactivator 1α (PGC-1α)-induced mitochondrial biogenesis. Proc. Natl Acad. Sci. USA 101, 6472–6477 (2004).
Cunningham, J. T. et al. mTOR controls mitochondrial oxidative function through a YY1-PGC-1α transcriptional complex. Nature 450, 736–740 (2007). This article describes mTORC1-dependent regulation of the transcriptional mitochondrial biogenesis programme by directing interaction of the transcription factor YY1 with PGC1α.
Puigserver, P. et al. Activation of PPARγ coactivator-1 through transcription factor docking. Science 286, 1368–1371 (1999).
Wallberg, A. E., Yamamura, S., Malik, S., Spiegelman, B. M. & Roeder, R. G. Coordination of p300-mediated chromatin remodeling and TRAP/mediator function through coactivator PGC-1α. Mol. Cell 12, 1137–1149 (2003).
Brandt, N., Dethlefsen, M. M., Bangsbo, J. & Pilegaard, H. PGC-1α and exercise intensity dependent adaptations in mouse skeletal muscle. PLoS ONE 12, e0185993 (2017).
Aguilo, F. et al. Deposition of 5-methylcytosine on enhancer RNAs enables the coactivator function of PGC-1α. Cell Rep. 14, 479–492 (2016).
Desautels, M. & Himms-Hagen, J. Parallel regression of cold-induced changes in ultrastructure, composition, and properties of brown adipose tissue mitochondria during recovery of rats from acclimation to cold. Can. J. Biochem. 58, 1057–1068 (1980).
Chouchani, E. T., Kazak, L. & Spiegelman, B. M. New advances in adaptive thermogenesis: UCP1 and beyond. Cell Metab. 29, 27–37 (2019).
Perry, C. G. R. & Hawley, J. A. Molecular basis of exercise-induced skeletal muscle mitochondrial biogenesis: historical advances, current knowledge, and future challenges. Cold Spring Harb. Perspect. Med. 8, a029686 (2018).
Liu, D. et al. Activation of mTORC1 is essential for β-adrenergic stimulation of adipose browning. J. Clin. Invest. 126, 1704–1716 (2016).
Mulligan, J. D., Gonzalez, A. A., Stewart, A. M., Carey, H. V. & Saupe, K. W. Upregulation of AMPK during cold exposure occurs via distinct mechanisms in brown and white adipose tissue of the mouse. J. Physiol. 580, 677–684 (2007).
Kato, H. et al. ER-resident sensor PERK is essential for mitochondrial thermogenesis in brown adipose tissue. Life Sci. Alliance 3, e201900576 (2020).
Lu, X. et al. Mitophagy controls beige adipocyte maintenance through a Parkin-dependent and UCP1-independent mechanism. Sci. Signal. 11, eaap8526 (2018).
Reznick, R. M. & Shulman, G. I. The role of AMP-activated protein kinase in mitochondrial biogenesis. J. Physiol. 574, 33–39 (2006).
Popov, D. V. Adaptation of skeletal muscles to contractile activity of varying duration and intensity: the role of PGC-1α. Biochemistry (Mosc.) 83, 613–628 (2018).
Spiegelman, B. Hormones, Metabolisme and the Benefits of Exercise. Trends in Molecular Medicine vol. 7 (Springer International Publishing, 2001).
Wu, H. et al. Regulation of mitochondrial biogenesis in skeletal muscle by caMK. Science 296, 349–352 (2002).
Gwinn, D. M. et al. AMPK phosphorylation of raptor mediates a metabolic checkpoint. Mol. Cell 30, 214–226 (2008).
Inoki, K., Zhu, T. & Guan, K. L. TSC2 mediates cellular energy response to control cell growth and survival. Cell 115, 577–590 (2003).
Sancak, Y. et al. The rag GTPases bind raptor and mediate amino acid signaling to mTORC1. Science 320, 1496–1501 (2008).
Sancak, Y. et al. Ragulator-Rag complex targets mTORC1 to the lysosomal surface and is necessary for its activation by amino acids. Cell 141, 290–303 (2010).
Nandagopal, N. & Roux, P. P. Regulation of global and specific mRNA translation by the mTOR signaling pathway. Translation 3, e983402 (2015).
Roux, P. P. & Topisirovic, I. Regulation of mRNA translation by signaling pathways. Cold Spring Harb. Perspect. Biol. 4, a012252 (2012).
Shirokikh, N. E. & Preiss, T. Translation initiation by cap-dependent ribosome recruitment: Recent insights and open questions. Wiley Interdiscip. Rev. RNA 9, e1473 (2018).
Hsieh, A. C. et al. The translational landscape of mTOR signalling steers cancer initiation and metastasis. Nature 485, 55–61 (2012).
Thoreen, C. C. et al. A unifying model for mTORC1-mediated regulation of mRNA translation. Nature 485, 109–113 (2012).
Morita, M. et al. MTORC1 controls mitochondrial activity and biogenesis through 4E-BP-dependent translational regulation. Cell Metab. 18, 698–711 (2013). This work discovers mTORC1 regulates the translation of numerous mitochondrial genes through phosphorylation of 4E-BP1/2, which promotes mitochondrial respiratory activity and ATP production.
Larsson, O. et al. Distinct perturbation of the translatome by the antidiabetic drug metformin. Proc. Natl Acad. Sci. USA 109, 8977–8982 (2012).
Gandin, V. et al. NanoCAGE reveals 5’ UTR features that define specific modes of translation of functionally related MTOR-sensitive mRNAs. Genome Res. 26, 636–648 (2016).
Elfakess, R. et al. Unique translation initiation of mRNAs-containing TISU element. Nucleic Acids Res. 39, 7598–7609 (2011).
Sinvani, H. et al. Translational tolerance of mitochondrial genes to metabolic energy stress involves TISU and eIF1-eIF4GI cooperation in start codon selection. Cell Metab. 21, 479–492 (2015). This article establishes that mitochondrial genes remain translationally active under nutrient deprivation, but not rapamycin treatment, owing to enrichment of TISU elements within their 5′ UTRs.
Elfakess, R. & Dikstein, R. A translation initiation element specific to mRNAs with very short 5′UTR that also regulates transcription. PLoS ONE 3, e3094 (2008).
Morita, M. et al. mTOR controls mitochondrial dynamics and cell survival via MTFP1. Mol. Cell 67, 922–935.e5 (2017).
Dumesic, P. A. et al. An evolutionarily conserved uORF regulates PGC1α and oxidative metabolism in mice, flies, and bluefin tuna. Cell Metab. 30, 190–200.e6 (2019).
McGillivray, P. et al. A comprehensive catalog of predicted functional upstream open reading frames in humans. Nucleic Acids Res. 46, 3326–3338 (2018).
Johnstone, T. G., Bazzini, A. A. & Giraldez, A. J. Upstream ORF s are prevalent translational repressors in vertebrates. EMBO J. 35, 706–723 (2016).
Lynch, M. R., Tran, M. T. & Parikh, S. M. PGC1α in the kidney. Am. J. Physiol. Ren. Physiol 314, F1–F8 (2018).
Han, S. H. et al. PGC-1a protects from notch-induced kidney fibrosis development. J. Am. Soc. Nephrol. 28, 3312–3322 (2017).
Pla-Martín, D. et al. CLUH granules coordinate translation of mitochondrial proteins with mTORC1 signaling and mitophagy. EMBO J. 39, e102731 (2020).
Gao, J. et al. CLUH regulates mitochondrial biogenesis by binding mRNAs of nuclear-encoded mitochondrial proteins. J. Cell Biol. 207, 213–223 (2014).
Schatton, D. et al. CLUH regulates mitochondrial metabolism by controlling translation and decay of target mRNAs. J. Cell Biol. 216, 675–693 (2017).
Sen, A. & Cox, R. T. Clueless is a conserved ribonucleoprotein that binds the ribosome at the mitochondrial outer membrane. Biol. Open 5, 195–203 (2016).
Sen, A., Kalvakuri, S., Bodmer, R. & Cox, R. T. Clueless, a protein required for mitochondrial function, interacts with the PINK1-Parkin complex in Drosophila. DMM Dis. Model. Mech. 8, 577–589 (2015).
Cannavino, J. et al. Regulation of cold-induced thermogenesis by the RNA binding protein FAM195A. Proc. Natl Acad. Sci. USA 118, e2104650118 (2021).
Molliex, A. et al. Phase separation by low complexity domains promotes stress granule assembly and drives pathological fibrillization. Cell 163, 123–133 (2015).
Kummer, E. & Ban, N. Mechanisms and regulation of protein synthesis in mitochondria. Nat. Rev. Mol. Cell Biol. 22, 307–325 (2021).
Montoya, J., Ojala, D. & Attardi, G. Distinctive features of the 5′-terminal sequences of the human mitochondrial mRNAs. Nature 290, 465–470 (1981).
Richman, T. R. et al. Loss of the RNA-binding protein TACO1 causes late-onset mitochondrial dysfunction in mice. Nat. Commun. 7, 11884 (2016).
Weraarpachai, W. et al. Mutation in TACO1, encoding a translational activator of COX I, results in cytochrome c oxidase deficiency and late-onset Leigh syndrome. Nat. Genet. 41, 833–837 (2009).
Taggart, J. C. & Li, G. W. Production of protein-complex components is stoichiometric and lacks general feedback regulation in eukaryotes. Cell Syst. 7, 580–589.e4 (2018).
Liu, L. et al. Nutrient sensing by the mitochondrial transcription machinery dictates oxidative phosphorylation. J. Clin. Invest. 124, 768–784 (2014).
Mukaneza, Y. et al. MTORC1 is required for expression of LRPPRC and cytochrome-c oxidase but not HIF-1α in leigh syndrome French Canadian type patient fibroblasts. Am. J. Physiol. Cell Physiol 317, C58–C67 (2019).
Weraarpachai, W. et al. Mutations in C12orf62, a factor that couples COX I synthesis with cytochrome c oxidase assembly, cause fatal neonatal lactic acidosis. Am. J. Hum. Genet. 90, 142–151 (2012).
Vance, J. E. MAM (mitochondria-associated membranes) in mammalian cells: lipids and beyond. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 1841, 595–609 (2014).
Hornbeck, P. V. et al. PhosphoSitePlus, 2014: mutations, PTMs and recalibrations. Nucleic Acids Res. 43, D512–D520 (2015).
Kampjut, D. & Sazanov, L. A. The coupling mechanism of mammalian respiratory complex I. Science 370, eabc4209 (2020). This is one of the most extensive high-resolution studies of the regulation and mechanism of action of respiratory complex I.
Friedkin, M. & Lehninger, A. L. Oxidation-coupled incorporation of inorganic radiophosphate into phospholipide and nucleic acid in a cell-free system. J. Biol. Chem. 177, 775–788 (1949).
Akabane, S. et al. PKA Regulates PINK1 stability and parkin recruitment to damaged mitochondria through phosphorylation of MIC60. Mol. Cell 62, 371–384 (2016).
Yadava, N., Potluri, P. & Scheffler, I. E. Investigations of the potential effects of phosphorylation of the MWFE and ESSS subunits on complex I activity and assembly. Int. J. Biochem. Cell Biol. 40, 447–460 (2008).
Rasmo, D. D. et al. Phosphorylation pattern of the NDUFS4 subunit of complex I of the mammalian respiratory chain. Mitochondrion 10, 464–471 (2010).
Samavati, L., Lee, I., Mathes, I., Lottspeich, F. & Hüttemann, M. Tumor necrosis factor α inhibits oxidative phosphorylation through tyrosine phosphorylation at subunit I of cytochrome c oxidase. J. Biol. Chem. 283, 21134–21144 (2008).
Acin-Perez, R., Gatti, D. L., Bai, Y. & Manfredi, G. Protein phosphorylation and prevention of cytochrome oxidase inhibition by ATP: coupled mechanisms of energy metabolism regulation. Cell Metab. 13, 712–719 (2011). This is the first and one of the most mechanistically detailed articles on the effect of PTMs on OXPHOS activity, together with Acin-Perez and Enriquez (2014).
De Rasmo, D., Panelli, D., Sardanelli, A. M. & Papa, S. cAMP-dependent protein kinase regulates the mitochondrial import of the nuclear encoded NDUFS4 subunit of complex I. Cell. Signal. 20, 989–997 (2008).
Srinivasan, S. et al. Oxidative stress induced mitochondrial protein kinase A mediates cytochrome C oxidase dysfunction. PLoS ONE 8, e77129 (2013).
Wang, Z. et al. Cyclin B1/Cdk1 coordinates mitochondrial respiration for cell-cycle G2/M progression. Dev. Cell 29, 217–232 (2014).
Morais, V. A. et al. PINK1 loss-of-function mutations affect mitochondrial complex I activity via NdufA10 ubiquinone uncoupling. Science 344, 203–207 (2014).
Ogura, M., Yamaki, J., Homma, M. K. & Homma, Y. Mitochondrial c-Src regulates cell survival through phosphorylation of respiratory chain components. Biochem. J. 447, 281–289 (2012).
Salvi, M., Morrice, N. A., Brunati, A. M. & Toninello, A. Identification of the flavoprotein of succinate dehydrogenase and aconitase as in vitro mitochondrial substrates of Fgr tyrosine kinase. FEBS Lett. 581, 5579–5585 (2007).
Acín-Pérez, R. et al. ROS-triggered phosphorylation of complex II by Fgr kinase regulates cellular adaptation to fuel use. Cell Metab. 19, 1020–1033 (2014).
Baeza, J., Smallegan, M. J. & Denu, J. M. Site-specific reactivity of nonenzymatic lysine acetylation. ACS Chem. Biol. 10, 122–128 (2015).
Narita, T., Weinert, B. T. & Choudhary, C. Functions and mechanisms of non-histone protein acetylation. Nat. Rev. Mol. Cell Biol. 20, 156–174 (2019).
Yang, W. et al. Mitochondrial sirtuin network reveals dynamic SIRT3-dependent deacetylation in response to membrane depolarization. Cell 167, 985–1000.e21 (2016).
Ahn, B. H. et al. A role for the mitochondrial deacetylase Sirt3 in regulating energy homeostasis. Proc. Natl Acad. Sci. USA 105, 14447–14452 (2008).
Porter, G. A., Urciuoli, W. R., Brookes, P. S. & Nadtochiy, S. M. SIRT3 deficiency exacerbates ischemia-reperfusion injury: implication for aged hearts. Am. J. Physiol. Hear. Circ. Physiol 306, H1602–H1609 (2014).
Koentges, C. et al. SIRT3 deficiency impairs mitochondrial and contractile function in the heart. Basic. Res. Cardiol. 110, 1–20 (2015).
Parodi-Rullán, R. M., Chapa-Dubocq, X., Rullán, P. J., Jang, S. & Javadov, S. High sensitivity of SIRT3 deficient hearts to ischemia-reperfusion is associated with mitochondrial abnormalities. Front. Pharmacol. 8, 275 (2017).
Finley, L. W. S. et al. Succinate dehydrogenase is a direct target of sirtuin 3 deacetylase activity. PLoS ONE 6, e23295 (2011).
Chen, Y. et al. Sirtuin-3 (SIRT3), a therapeutic target with oncogenic and tumor-suppressive function in cancer. Cell Death Dis 5, e1047 (2014).
Xiao, H. et al. A quantitative tissue-specific landscape of protein redox regulation during aging. Cell 180, 968–983.e24 (2020).
Mills, E. L. et al. Cysteine 253 of UCP1 regulates energy expenditure and sex-dependent adipose tissue inflammation. Cell Metab. 34, 140–157.e8 (2022).
Chouchani, E. T. et al. Cardioprotection by S-nitrosation of a cysteine switch on mitochondrial complex I. Nat. Med. 19, 753–759 (2013).
Yin, Z. et al. Structural basis for a complex I mutation that blocks pathological ROS production. Nat. Commun. 12, 707 (2021).
Burger, N. et al. ND3 Cys39 in complex I is exposed during mitochondrial respiration. Cell Chem. Biol. 29, 636–649.e14 (2022).
Araiso, Y. et al. Structure of the mitochondrial import gate reveals distinct preprotein paths. Nature 575, 395–401 (2019).
Tucker, K. & Park, E. Cryo-EM structure of the mitochondrial protein-import channel TOM complex at near-atomic resolution. Nat. Struct. Mol. Biol. 26, 1158–1166 (2019).
Roise, D. et al. Amphiphilicity is essential for mitochondrial presequence function. EMBO J. 7, 649–653 (1988).
Allison, D. S. & Schatz, G. Artificial mitochondrial presequences. Proc. Natl Acad. Sci. USA 83, 9011–9015 (1986).
Schmidt, O. et al. Regulation of mitochondrial protein import by cytosolic kinases. Cell 144, 227–239 (2011). This is the first report on the effect of PTMs on import machinery function.
Pfanner, N. & Neupert, W. Distinct steps in the import of ADP/ATP carrier into mitochondria. J. Biol. Chem. 262, 7528–7536 (1987).
Pfanner, N., Tropschug, M. & Neupert, W. Mitochondrial protein import: nucleoside triphosphates are involved in conferring import-competence to precursors. Cell 49, 815–823 (1987).
Young, J. C., Hoogenraad, N. J. & Hartl, F. U. Molecular chaperones Hsp90 and Hsp70 deliver preproteins to the mitochondrial import receptor Tom70. Cell 112, 41–50 (2003). This work provides the first evidence connecting cytosolic chaperones with the import of mitochondrial proteins through the receptor TOM70.
Bhangoo, M. K. et al. Multiple 40-kDa heat-shock protein chaperones function in Tom70-dependent mitochondrial import. Mol. Biol. Cell 18, 3414–3428 (2007).
Rampelt, H. et al. The mitochondrial carrier pathway transports non-canonical substrates with an odd number of transmembrane segments. BMC Biol. 18, 2 (2020).
Yamamoto, H. et al. Roles of Tom70 in import of presequence-containing mitochondrial proteins. J. Biol. Chem. 284, 31635–31646 (2009).
Backes, S. et al. Tom70 enhances mitochondrial preprotein import efficiency by binding to internal targeting sequences. J. Cell Biol. 217, 1369–1382 (2018).
Fan, A. C. Y. et al. Interaction between the human mitochondrial import receptors Tom20 and Tom70 in vitro suggests a chaperone displacement mechanism. J. Biol. Chem. 286, 32208–32219 (2011).
Wei, X. et al. Mutations in TOMM70 lead to multi-OXPHOS deficiencies and cause severe anemia, lactic acidosis, and developmental delay. J. Hum. Genet. 65, 231–240 (2020).
Brix, J., Dietmeier, K. & Pfanner, N. Differential recognition of preproteins by the purified cytosolic domains of the mitochondrial import receptors Tom20, Tom22, and Tom70. J. Biol. Chem. 272, 20730–20735 (1997).
Kreimendahl, S., Schwichtenberg, J., Günnewig, K., Brandherm, L. & Rassow, J. The selectivity filter of the mitochondrial protein import machinery. BMC Biol. 18, 1–23 (2020).
Tripathi, A., Mandon, E. C., Gilmore, R. & Rapoport, T. A. Two alternative binding mechanisms connect the protein translocation Sec71-Sec72 complex with heat shock proteins. J. Biol. Chem. 292, 8007–8018 (2017).
Gordon, D. E. et al. Comparative host-coronavirus protein interaction networks reveal pan-viral disease mechanisms. Science 370, eabe9403 (2020).
Liu, X. Y., Wei, B., Shi, H. X., Shan, Y. F. & Wang, C. Tom70 mediates activation of interferon regulatory factor 3 on mitochondria. Cell Res. 20, 994–1011 (2010).
Filadi, R. et al. TOM70 sustains cell bioenergetics by promoting IP3R3-mediated ER to mitochondria Ca2+transfer. Curr. Biol. 28, 369–382.e6 (2018).
An, Y. A. et al. Dysregulation of amyloid precursor protein impairs adipose tissue mitochondrial function and promotes obesity. Nat. Metab. 1, 1243–1257 (2019).
Leney, A. C., El Atmioui, D., Wu, W., Ovaa, H. & Heck, A. J. R. Elucidating crosstalk mechanisms between phosphorylation and O-GlcNAcylation. Proc. Natl Acad. Sci. USA 114, E7255–E7261 (2017).
Tarrant, M. K. et al. Regulation of CK2 by phosphorylation and O-GlcNAcylation revealed by semisynthesis. Nat. Chem. Biol. 8, 262–269 (2012).
Shinoda, K. et al. Phosphoproteomics identifies CK2 as a negative regulator of beige adipocyte thermogenesis and energy expenditure. Cell Metab. 22, 997–1008 (2015).
Manni, S. et al. Protein kinase CK2 protects multiple myeloma cells from ER stress-induced apoptosis and from the cytotoxic effect of HSP90 inhibition through regulation of the unfolded protein response. Clin. Cancer Res. 18, 1888–1900 (2012).
Hessenauer, A., Schneider, C. C., Götz Claudia, C. & Montenarh, M. CK2 inhibition induces apoptosis via the ER stress response. Cell. Signal. 23, 145–151 (2011).
Voos, W. Chaperone-protease networks in mitochondrial protein homeostasis. Biochim. Biophys. Acta Mol. Cell Res. 1833, 388–399 (2013).
Matsushima, Y. et al. Mitochondrial Lon protease is a gatekeeper for proteins newly imported into the matrix. Commun. Biol. 4, 974 (2021).
Shin, C. S. et al. LONP1 and mtHSP70 cooperate to promote mitochondrial protein folding. Nat. Commun. 12, 265 (2021).
Rep, M. et al. Promotion of mitochondrial membrane complex assembly by a proteolytically inactive yeast Lon. Science 274, 103–106 (1996).
Ghosh, J. C. et al. Akt phosphorylation of mitochondrial Lonp1 protease enables oxidative metabolism and advanced tumor traits. Oncogene 38, 6926–6939 (2019).
MacVicar, T. et al. Lipid signalling drives proteolytic rewiring of mitochondria by YME1L. Nature 575, 361–365 (2019).
Pfanner, N., Warscheid, B. & Wiedemann, N. Mitochondrial proteins: from biogenesis to functional networks. Nat. Rev. Mol. Cell Biol. 20, 267–284 (2019). This is one of the most compelling reviews that discusses the mechanisms and roles of import machineries in organizing mitochondrial function.
Priesnitz, C., Pfanner, N. & Becker, T. Studying Protein Import into Mitochondria. Methods in Cell Biology vol. 155 (Elsevier, 2020).
Guerrero-Castillo, S. et al. The assembly pathway of mitochondrial respiratory chain complex I. Cell Metab. 25, 128–139 (2017).
Vercellino, I. & Sazanov, L. A. The assembly, regulation and function of the mitochondrial respiratory chain. Nat. Rev. Mol. Cell Biol. 23, 141–161 (2022).
Schägger, H. & Pfeiffer, K. The ratio of oxidative phosphorylation complexes I-V in Bovine heart mitochondria and the composition of respiratory chain supercomplexes. J. Biol. Chem. 276, 37861–37867 (2001).
Greggio, C. et al. Enhanced respiratory chain supercomplex formation in response to exercise in human skeletal muscle. Cell Metab. 25, 301–311 (2017).
Schägger, H. & Pfeiffer, K. Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J. 19, 1777–1783 (2000). This article overturns the paradigm on respiratory chain complex organization. Using blue native PAGE, the authors establish that yeast and mammalian respiratory chain complexes superassemble.
Acin-Perez, R. & Enriquez, J. A. The function of the respiratory supercomplexes: the plasticity model. Biochim. Biophys. Acta Bioenerg. 1837, 444–450 (2014).
Acín-Pérez, R., Fernández-Silva, P., Peleato, M. L., Pérez-Martos, A. & Enriquez, J. A. Respiratory active mitochondrial supercomplexes. Mol. Cell 32, 529–539 (2008).
Gonzalez-Franquesa, A. et al. Mass-spectrometry-based proteomics reveals mitochondrial supercomplexome plasticity. Cell Rep. 35, 109180 (2021).
Lapuente-Brun, E. et al. Supercomplex assembly determines electron flux in the mitochondrial electron transport chain. Science 340, 1567–1570 (2013).
Lopez-Fabuel, I. et al. Complex I assembly into supercomplexes determines differential mitochondrial ROS production in neurons and astrocytes. Proc. Natl Acad. Sci. USA 113, 13063–13068 (2016).
Calvo, E. et al. Functional role of respiratory supercomplexes in mice: SCAF1 relevance and segmentation of the Qpool. Sci. Adv. 6, eaba7509 (2020).
Vercellino, I. & Sazanov, L. A. Structure and assembly of the mammalian mitochondrial supercomplex CIII2CIV. Nature 598, 364–367 (2021). This article provides the first published structure of mammalian supercomplex III2 + IV and characterization of SCAF1-dependent superassembly and kinetic activity.
Moe, A., Trani, J. D., Rubinstein, J. L. & Brzezinski, P. Cryo-EM structure and kinetics reveal electron transfer by 2D diffusion of cytochrome c in the yeast III-IV respiratory supercomplex. Proc. Natl Acad. Sci. USA 118, e2021157118 (2021).
Berndtsson, J. et al. Respiratory supercomplexes enhance electron transport by decreasing cytochrome c diffusion distance. EMBO Rep 21, e51015 (2020).
Blanchi, C., Genova, M. L., Castelli, G. P. & Lenaz, G. The mitochondrial respiratory chain is partially organized in a supercomplex assembly: kinetic evidence using flux control analysis. J. Biol. Chem. 279, 36562–36569 (2004).
Acín-Pérez, R. et al. Respiratory complex III is required to maintain complex I in mammalian mitochondria. Mol. Cell 13, 805–815 (2004).
Calvaruso, M. A. et al. Mitochondrial complex III stabilizes complex I in the absence of NDUFS4 to provide partial activity. Hum. Mol. Genet. 21, 115–120 (2012).
Schägger, H. et al. Significance of respirasomes for the assembly/stability of human respiratory chain complex I. J. Biol. Chem. 279, 36349–36353 (2004).
Protasoni, M. et al. Respiratory supercomplexes act as a platform for complex III -mediated maturation of human mitochondrial complexes I and IV. EMBO J. 39, e102817 (2020).
Blaza, J. N., Serreli, R., Jones, A. J. Y., Mohammed, K. & Hirst, J. Kinetic evidence against partitioning of the ubiquinone pool and the catalytic relevance of respiratory-chain supercomplexes. Proc. Natl Acad. Sci. USA 111, 15735–15740 (2014).
Lobo-Jarne, T. et al. Human COX7A2L regulates complex III biogenesis and promotes supercomplex organization remodeling without affecting mitochondrial bioenergetics. Cell Rep. 25, 1786–1799.e4 (2018).
Cogliati, S. et al. Mechanism of super-assembly of respiratory complexes III and IV. Nature 539, 579–582 (2016).
Pérez-Pérez, R. et al. COX7A2L is a mitochondrial complex III binding protein that stabilizes the III2+IV supercomplex without affecting respirasome formation. Cell Rep. 16, 2387–2398 (2016).
Ikeda, K. et al. Mitochondrial supercomplex assembly promotes breast and endometrial tumorigenesis by metabolic alterations and enhanced hypoxia tolerance. Nat. Commun. 10, 4108 (2019).
Hollinshead, K. E. R. et al. Respiratory supercomplexes promote mitochondrial efficiency and growth in severely hypoxic pancreatic cancer. Cell Rep. 33, 108231 (2020).
Mirali, S. et al. The mitochondrial peptidase, neurolysin, regulates respiratory chain supercomplex formation and is necessary for AML viability. Sci. Transl. Med. 12, eaaz8264 (2020).
García-Poyatos, C. et al. Scaf1 promotes respiratory supercomplexes and metabolic efficiency in zebrafish. EMBO Rep. 21, e50287 (2020).
Chan, D. C. Mitochondrial dynamics and its involvement in disease. Annu. Rev. Pathol. Mech. Dis. 15, 235–259 (2020).
Friedman, J. R. & Nunnari, J. Mitochondrial form and function. Nature 505, 335–343 (2014).
Friedman, J. R. et al. ER tubules mark sites of mitochondrial division. Science 334, 358–362 (2011).
Rambold, A. S., Kostelecky, B., Elia, N. & Lippincott-Schwartz, J. Tubular network formation protects mitochondria from autophagosomal degradation during nutrient starvation. Proc. Natl Acad. Sci. USA 108, 10190–10195 (2011).
Gomes, L. C., Benedetto, G. D. & Scorrano, L. During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598 (2011).
Kim, H. et al. Fine-tuning of Drp1/Fis1 availability by AKAP121/Siah2 regulates mitochondrial adaptation to hypoxia. Mol. Cell 44, 532–544 (2011).
Yao, C. H. et al. Mitochondrial fusion supports increased oxidative phosphorylation during cell proliferation. eLife 8, e41351 (2019).
Wikstrom, J. D. et al. Hormone-induced mitochondrial fission is utilized by brown adipocytes as an amplification pathway for energy expenditure. EMBO J. 33, 418–436 (2014).
Coronado, M. et al. Physiological mitochondrial fragmentation is a normal cardiac adaptation to increased energy demand. Circ. Res. 122, 282–295 (2018).
Waters, L. R., Ahsan, F. M., Wolf, D. M., Shirihai, O. & Teitell, M. A. Initial B cell activation induces metabolic reprogramming and mitochondrial remodeling. iScience 5, 99–109 (2018).
Ron-Harel, N. et al. Mitochondrial biogenesis and proteome remodeling promote one-carbon metabolism for T cell activation. Cell Metab. 24, 104–117 (2016).
Cogliati, S. et al. Mitochondrial cristae shape determines respiratory chain supercomplexes assembly and respiratory efficiency. Cell 155, 160–171 (2013). This article establishes the connection between cristae morphology and respiratory chain complex assembly and function.
Bal, N. C. et al. Both brown adipose tissue and skeletal muscle thermogenesis processes are activated during mild to severe cold adaptation in mice. J. Biol. Chem. 292, 16616–16625 (2017).
Kondadi, A. K. et al. Cristae undergo continuous cycles of fusion and fission in a MICOS-dependent manner. Preprint at bioRxiv https://doi.org/10.1101/654541 (2019).
Patten, D. A. et al. OPA1-dependent cristae modulation is essential for cellular adaptation to metabolic demand. EMBO J. 33, 2676–2691 (2014).
Tang, J. et al. Sam50–Mic19–Mic60 axis determines mitochondrial cristae architecture by mediating mitochondrial outer and inner membrane contact. Cell Death Differ. 27, 146–160 (2020).
Ott, C., Dorsch, E., Fraunholz, M., Straub, S. & Kozjak-Pavlovic, V. Detailed analysis of the human mitochondrial contact site complex indicate a hierarchy of subunits. PLoS ONE 10, 1–15 (2015).
Li, H. et al. Mic60/Mitofilin determines MICOS assembly essential for mitochondrial dynamics and mtDNA nucleoid organization. Cell Death Differ. 23, 380–392 (2016).
Friedman, J. R., Mourier, A., Yamada, J., Michael McCaffery, J. & Nunnari, J. MICOS coordinates with respiratory complexes and lipids to establish mitochondrial inner membrane architecture. eLife 2015, 1–61 (2015).
Strauss, M., Hofhaus, G., Schröder, R. R. & Kühlbrandt, W. Dimer ribbons of ATP synthase shape the inner mitochondrial membrane. EMBO J. 27, 1154–1160 (2008).
Daum, B., Walter, A., Horst, A., Osiewacz, H. D. & Kühlbrandt, W. Age-dependent dissociation of ATP synthase dimers and loss of inner-membrane cristae in mitochondria. Proc. Natl Acad. Sci. USA 110, 15301–15306 (2013).
Paumard, P. The ATP synthase is involved in generating mitochondrial cristae morphology. EMBO J. 21, 221–230 (2002).
Blum, T. B., Hahn, A., Meier, T., Davies, K. M. & Kühlbrandt, W. Dimers of mitochondrial ATP synthase induce membrane curvature and self-assemble into rows. Proc. Natl Acad. Sci. USA 116, 4250–4255 (2019).
Anand, R. et al. MIC26 and MIC27 cooperate to regulate cardiolipin levels and the landscape of OXPHOS complexes. Life Sci. Alliance 3, 1–17 (2020).
Pfeiffer, K. et al. Cardiolipin stabilizes respiratory chain supercomplexes. J. Biol. Chem. 278, 52873–52880 (2003).
Kramarova, T. V. et al. Mitochondrial ATP synthase levels in brown adipose tissue are governed by the c-Fo subunit P1 isoform. FASEB J. 22, 55–63 (2008).
Rampelt, H., Zerbes, R. M., van der Laan, M. & Pfanner, N. Role of the mitochondrial contact site and cristae organizing system in membrane architecture and dynamics. Biochim. Biophys. Acta Mol. Cell Res. 1864, 737–746 (2017).
Banci, L. et al. Structural characterization of CHCHD5 and CHCHD7: two atypical human twin CX9C proteins. J. Struct. Biol. 180, 190–200 (2012).
Sakowska, P. et al. The oxidation status of Mic19 regulates MICOS assembly. Mol. Cell. Biol. 35, 4222–4237 (2015).
Zhou, Z. D., Saw, W. T. & Tan, E. K. Mitochondrial CHCHD-containing proteins: physiologic functions and link with neurodegenerative diseases. Mol. Neurobiol. 54, 5534–5546 (2017).
Ueda, E. et al. Myristoyl group-aided protein import into the mitochondrial intermembrane space. Sci. Rep. 9, 1185 (2019).
Tsai, P. I. et al. PINK1 Phosphorylates MIC60/mitofilin to control structural plasticity of mitochondrial crista junctions. Mol. Cell 69, 744–756.e6 (2018).
Kawano, S. et al. Structure-function insights into direct lipid transfer between membranes by Mmm 1-Mdm 12 of ERMES. J. Cell Biol. 217, 959–974 (2018).
Guillén-Samander, A. et al. VPS13D bridges the ER to mitochondria and peroxisomes via Miro. J. Cell Biol. 220, e202010004 (2021).
Koob, S., Barrera, M., Anand, R. & Reichert, A. S. The non-glycosylated isoform of MIC26 is a constituent of the mammalian MICOS complex and promotes formation of crista junctions. Biochim. Biophys. Acta Mol. Cell Res. 1853, 1551–1563 (2015).
Van Den Brink-Van Der Laan, E., Antoinette Killian, J. & De Kruijff, B. Nonbilayer lipids affect peripheral and integral membrane proteins via changes in the lateral pressure profile. Biochim. Biophys. Acta Biomembr. 1666, 275–288 (2004).
Mejia, E. M. & Hatch, G. M. Mitochondrial phospholipids: role in mitochondrial function. J. Bioenerg. Biomembr. 48, 99–112 (2016).
Osman, C., Voelker, D. R. & Langer, T. Making heads or tails of phospholipids in mitochondria. J. Cell Biol. 192, 7–16 (2011).
Böttinger, L. et al. Phosphatidylethanolamine and cardiolipin differentially affect the stability of mitochondrial respiratory chain supercomplexes. J. Mol. Biol. 423, 677–686 (2012).
Tasseva, G. et al. Phosphatidylethanolamine deficiency in mammalian mitochondria impairs oxidative phosphorylation and alters mitochondrial morphology. J. Biol. Chem. 288, 4158–4173 (2013).
Das, S. et al. ATP citrate lyase improves mitochondrial function in skeletal muscle. Cell Metab. 21, 868–876 (2015).
Lynes, M. D. et al. Cold-activated lipid dynamics in adipose tissue highlights a role for cardiolipin in thermogenic metabolism. Cell Rep. 24, 781–790 (2018).
May, F. J. et al. Lipidomic adaptations in white and brown adipose tissue in response to exercise demonstrate molecular species-specific remodeling. Cell Rep. 18, 1558–1572 (2017).
Marcher, A. B. et al. RNA-seq and mass-spectrometry-based lipidomics reveal extensive changes of glycerolipid pathways in brown adipose tissue in response to cold. Cell Rep. 13, 2000–2013 (2015).
Jain, I. H. et al. Genetic screen for cell fitness in high or low oxygen highlights mitochondrial and lipid metabolism. Cell 181, 716–727.e11 (2020).
Baker, C. D., Ball, W. B., Pryce, E. N. & Gohil, V. M. Specific requirements of nonbilayer phospholipids in mitochondrial respiratory chain function and formation. Mol. Biol. Cell 27, 2161–2171 (2016).
Tuller, G., Nemec, T., Hrastnik, C. & Daum, G. Lipid composition of subcellular membranes of an FY1679-derived haploid yeast wild-type strain grown on different carbon sources. Yeast 15, 1555–1564 (1999).
Yan, F., Zhao, H. & Zeng, Y. Lipidomics: a promising cancer biomarker. Clin. Transl. Med. 7, 21 (2018).
Dean, J. M. & Lodhi, I. J. Structural and functional roles of ether lipids. Protein Cell 9, 196–206 (2018).
Park, H. et al. Peroxisome-derived lipids regulate adipose thermogenesis by mediating cold-induced mitochondrial fission. J. Clin. Invest. 129, 694–711 (2019).
Kimura, T. et al. Substantial decrease in plasmalogen in the heart associated with tafazzin deficiency. Biochemistry 57, 2162–2175 (2018).
Kimura, T. et al. Plasmalogen loss caused by remodeling deficiency in mitochondria. Life Sci. Alliance 2, e201900348 (2019).
Zhu, Y. et al. Alkylglyceronephosphate synthase (AGPS) alters lipid signaling pathways and supports chemotherapy resistance of glioma and hepatic carcinoma cell lines. Asian Pac. J. Cancer Prev. 15, 3219–3226 (2014).
Benjamin, D. I. et al. Ether lipid generating enzyme AGPS alters the balance of structural and signaling lipids to fuel cancer pathogenicity. Proc. Natl Acad. Sci. USA 110, 14912–14917 (2013).
Messias, M. C. F., Mecatti, G. C., Priolli, D. G. & De Oliveira Carvalho, P. Plasmalogen lipids: Functional mechanism and their involvement in gastrointestinal cancer. Lipids Health Dis. 17, 41 (2018).
Wu, M., Gu, J., Guo, R., Huang, Y. & Yang, M. Structure of mammalian respiratory supercomplex I1III2IV1. Cell 167, 1598–1609.e10 (2016).
Sun, F. et al. Crystal structure of mitochondrial respiratory membrane protein complex II. Cell 121, 1043–1057 (2005).
Spikes, T. E., Montgomery, M. G. & Walker, J. E. Structure of the dimeric ATP synthase from bovine mitochondria. Proc. Natl Acad. Sci. USA 117, 23519–23526 (2020).
Pebay-Peyroula, E. et al. Structure of mitochondrial ADP/ATP carrier in complex with carboxyatractyloside. Nature 426, 39–44 (2003).
Berardi, M. J., Shih, W. M., Harrison, S. C. & Chou, J. J. Mitochondrial uncoupling protein 2 structure determined by NMR molecular fragment searching. Nature 476, 109–113 (2011).
Park, Y., Reyna-Neyra, A., Philippe, L. & Thoreen, C. C. mTORC1 balances cellular amino acid supply with demand for protein synthesis through post-transcriptional control of ATF4. Cell Rep. 19, 1083–1090 (2017).
Davies, K. M., Anselmi, C., Wittig, I., Faraldo-Gómez, J. D. & Kühlbrandt, W. Structure of the yeast F1Fo-ATP synthase dimer and its role in shaping the mitochondrial cristae. Proc. Natl Acad. Sci. USA 109, 13602–13607 (2012).
Nicholls, D. G. & Ferguson, S. J. Respiratory chains. in Bioenergetics 91–157 (Elsevier, 2013).
Guo, R., Gu, J., Zong, S., Wu, M. & Yang, M. Structure and mechanism of mitochondrial electron transport chain. Biomed. J. 41, 9–20 (2018).
Letts, J. A. & Sazanov, L. A. Clarifying the supercomplex: the higher-order organization of the mitochondrial electron transport chain. Nat. Struct. Mol. Biol. 24, 800–808 (2017).
Sousa, J. S., Mills, D. J., Vonck, J. & Kühlbrandt, W. Functional asymmetry and electron flow in the bovine respirasome. eLife 5, e21290 (2016).
Gu, J. et al. The architecture of the mammalian respirasome. Nature 537, 639–643 (2016).
Letts, J. A., Fiedorczuk, K. & Sazanov, L. A. The architecture of respiratory supercomplexes. Nature 537, 644–648 (2016).
Martínez-Reyes, I. & Chandel, N. S. Mitochondrial TCA cycle metabolites control physiology and disease. Nat. Commun. 11, 102 (2020).
Arnold, S. & Kadenbach, B. Cell respiration is controlled by ATP, an allosteric inhibitor of cytochrome-c oxidase. Eur. J. Biochem. 249, 350–354 (1997).
Schneeberger, M. et al. Mitofusin 2 in POMC neurons connects ER stress with leptin resistance and energy imbalance. Cell 155, 172–187 (2013).
Tubbs, E. et al. Mitochondria-associated endoplasmic reticulum membrane (MAM) integrity is required for insulin signaling and is implicated in hepatic insulin resistance. Diabetes 63, 3279–3294 (2014).
De Brito, O. M. & Scorrano, L. Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature 456, 605–610 (2008).
Muñoz, J. P. et al. Mfn2 modulates the UPR and mitochondrial function via repression of PERK. EMBO J. 32, 2348–2361 (2013).
Abrisch, R. G., Gumbin, S. C., Wisniewski, B. T., Lackner, L. L. & Voeltz, G. K. Fission and fusion machineries converge at ER contact sites to regulate mitochondrial morphology. J. Cell Biol. 219, e201911122 (2020).
Bravo, R. et al. Increased ER-mitochondrial coupling promotes mitochondrial respiration and bioenergetics during early phases of ER stress. J. Cell Sci. 124, 2143–2152 (2011).
Basso, V., Marchesan, E. & Ziviani, E. A trio has turned into a quartet: DJ-1 interacts with the IP3R-Grp75-VDAC complex to control ER-mitochondria interaction. Cell Calcium 87, 102186 (2020).
De vos, K. J. et al. VAPB interacts with the mitochondrial protein PTPIP51 to regulate calcium homeostasis. Hum. Mol. Genet. 21, 1299–1311 (2012).
Stoica, R. et al. ER-mitochondria associations are regulated by the VAPB-PTPIP51 interaction and are disrupted by ALS/FTD-associated TDP-43. Nat. Commun. 5, 3996 (2014).
Stoica, R. et al. ALS/FTD-associated FUS activates GSK-3β to disrupt the VAPB–PTPIP51 interaction and ER–mitochondria associations. EMBO Rep. 17, 1326–1342 (2016).
Naon, D. et al. Critical reappraisal confirms that mitofusin 2 is an endoplasmic reticulum-mitochondria tether. Proc. Natl Acad. Sci. USA 113, 11249–11254 (2016).
Mancuso, M. et al. Fatigue and exercise intolerance in mitochondrial diseases. Literature revision and experience of the Italian Network of Mitochondrial Diseases. Neuromuscul. Disord. 22, S226–S229 (2012).
Mito, T. et al. Mosaic dysfunction of mitophagy in mitochondrial muscle disease. Cell Metab. 34, 197–208.e5 (2022).
Kleiner, S. et al. Development of insulin resistance in mice lacking PGC-1α in adipose tissues. Proc. Natl Acad. Sci. USA 109, 9635–9640 (2012).
Cohen, P. et al. Ablation of PRDM16 and beige adipose causes metabolic dysfunction and a subcutaneous to visceral fat switch. Cell 156, 304–316 (2014).
Lehman, J. J. & Kelly, D. P. Transcriptional activation of energy metabolic switches in the developing and hypertrophied heart. Clin. Exp. Pharmacol. Physiol. 29, 339–345 (2002).
Chambers, J. M. & Wingert, R. A. PGC-1α in disease: recent renal insights into a versatile metabolic regulator. Cells 9, 2234 (2020).
Zhang, L., Liu, J., Zhou, F., Wang, W. & Chen, N. PGC-1α ameliorates kidney fibrosis in mice with diabetic kidney disease through an antioxidative mechanism. Mol. Med. Rep. 17, 4490–4498 (2018).
Piccinin, E. et al. PGC-1s in the spotlight with Parkinson’s disease. Int. J. Mol. Sci. 22, 3487 (2021).
Kumar, M. et al. Defects in mitochondrial biogenesis drive mitochondrial alterations in PARKIN-deficient human dopamine neurons. Stem Cell Rep. 15, 629–645 (2020).
Lv, J. et al. PGC-1α sparks the fire of neuroprotection against neurodegenerative disorders. Ageing Res. Rev. 44, 8–21 (2018).
Dumauthioz, N. et al. Enforced PGC-1α expression promotes CD8 T cell fitness, memory formation and antitumor immunity. Cell. Mol. Immunol. 18, 1761–1771 (2021).
Gerbec, Z. J. et al. Conditional deletion of PGC-1α results in energetic and functional defects in NK cells. iScience 23, 101454 (2020).
Bertero, E., Kutschka, I., Maack, C. & Dudek, J. Cardiolipin remodeling in Barth syndrome and other hereditary cardiomyopathies. Biochim. Biophys. Acta Mol. Basis Dis. 1866, 165803 (2020).
MacKenzie, J. A. & Payne, R. M. Mitochondrial protein import and human health and disease. Biochim. Biophys. Acta Mol. Basis Dis. 1772, 509–523 (2007).
This work was partially supported by grants from the US National Institutes of Health R01 DK089883 (NIDDK), R01 DK081418 (NIDDK), R01 DK117655 (NIDDK), R01 CA181217 (NCI), 9R56 AG074527 (NIA), R01 GM121452 (NIGMS), and the Claudia Adams Barr Award to P.P. F32 GM125243 (NIGMS) and the Charles A. King Trust Postdoctoral Fellowship Program were awarded to C.F.B. P.L.-M. is supported by the Human Frontier Science Program (LT-000033/2019-L).
The authors declare no competing interests.
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Invaginations of the inner mitochondrial membrane that increase the surface area of respiratory reactions and harbour the respiratory complexes.
- Brown and beige adipose tissues
Thermogenic fat cells with abundant mitochondria that oxidize glucose, fatty acids and branched-chain amino acids to generate heat.
- Uncoupling protein 1
(UCP1). Inner mitochondrial membrane protein expressed in thermogenic tissues that dissipates membrane potential from the respiratory complexes and sustains thermogenesis.
Porphyrin group coordinating an iron atom required for electron transfer or oxygen transport.
- Iron–sulfur clusters
Iron sulfide molecules that transfer electrons across respiratory complexes.
- ER stress
A condition that occurs when the capacity of the endoplasmic reticulum (ER) lumen to fold proteins is saturated. ER stress transduces signals to other organelles such as mitochondria to adapt cellular metabolism to satisfy energy demands.
- Mediator complex
A multiprotein complex that transduces signals from transcription factors to RNA polymerase II to control gene expression.
- Nuclear receptors
Ligand-regulated transcription factors that are activated by steroid hormones and other lipid-related molecules.
A group of chemical neurotransmitters, including dopamine, adrenaline and noradrenaline, that are released into the blood upon stress and modulate beige and brown fat tissue activity.
Mammalian target of rapamycin, a nutrient-sensor kinase involved in the control of cellular growth, survival, metabolism and immunity.
AMP-activated protein kinase that senses fluctuations in the ATP/AMP ratio.
Also known as protein kinase B (PKB), a group of serine/threonine kinases that respond to a myriad of external stimuli and include AKT1, AKT2 and AKT3.
- Translation initiator of short 5′ UTR (TISU) elements
Sequence elements that are downstream of transcription start sites and regulate both transcriptional and translational initiation and are present in mRNAs with very short 5′ untranslated regions (UTRs).
- PINK1–parkin mitophagy pathway
A quality control pathway that marks damaged mitochondria to promote their autophagy-mediated destruction.
- Branched-chain amino acids
Amino acids, including valine, leucine and isoleucine, that can be oxidized in the cell to obtain energy.
- Unfolded protein response
A process that regulates a transcriptional and translational response to endoplasmic reticulum protein folding stress.
- Internal mitochondrial targeting sequence-like signals
Peptide sequences present within proteins destined for mitochondria that interact with import receptors and increase import competence.
- SEC translocon
Protein complex embedded in the endoplasmic reticulum membrane that transports proteins from and through the endoplasmic reticulum lumen.
- ADP/ATP carrier
An inner mitochondrial membrane transporter that exchanges ATP/ADP.
- Phosphate carrier
An inner mitochondrial membrane transporter of phosphate.
- AAA+ proteases
Subset of ATPase proteases that participate in diverse quality control mechanisms in mitochondria and the cytosol (26S proteasome).
- Substrate channelling
Biochemical phenomenon whereby the intermediate product from one enzyme is shuttled as a substrate to the next enzyme before equilibration with the bulk aqueous solvent. This model is contested for mitochondrial respiratory supercomplexes, but refers to the presence of functionally distinct pools of ubiquinone or cytochrome c within supercomplexess that pass between complex I and complex III2 or complex III2 and complex IV, respectively.
A class of phospholipids containing a sphingosine backbone. Sphingolipids facilitate mitochondrial function and cellular signalling responses, but their accumulation correlates with mitochondrial dysfunction and chronic metabolic diseases such as type 2 diabetes.
- sn-1 position
The first stereochemical position on a glycerol moiety to which a fatty acid is attached.
- Barth syndrome
A rare X-linked genetic disorder of cardiolipin metabolism that presents with cardiomyopathy, neutropenia and muscle weakness.
- Substantia nigra
Basal ganglia structure in the brain that plays important roles in behaviour–reward neuronal programmes.
- Sengers syndrome
A rare autosomal condition characterized by myocardiopathy, lactic acidosis, muscle weakness and short life expectancy.
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Bennett, C.F., Latorre-Muro, P. & Puigserver, P. Mechanisms of mitochondrial respiratory adaptation. Nat Rev Mol Cell Biol 23, 817–835 (2022). https://doi.org/10.1038/s41580-022-00506-6
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