Owing to their ability to efficiently generate ATP required to sustain normal cell function, mitochondria are often considered the ‘powerhouses of the cell’. However, our understanding of the role of mitochondria in cell biology recently expanded when we recognized that they are key platforms for a plethora of cell signalling cascades. This functional versatility is tightly coupled to constant reshaping of the cellular mitochondrial network in a series of processes, collectively referred to as mitochondrial membrane dynamics and involving organelle fusion and fission (division) as well as ultrastructural remodelling of the membrane. Accordingly, mitochondrial dynamics influence and often orchestrate not only metabolism but also complex cell signalling events, such as those involved in regulating cell pluripotency, division, differentiation, senescence and death. Reciprocally, mitochondrial membrane dynamics are extensively regulated by post-translational modifications of its machinery and by the formation of membrane contact sites between mitochondria and other organelles, both of which have the capacity to integrate inputs from various pathways. Here, we discuss mitochondrial membrane dynamics and their regulation and describe how bioenergetics and cellular signalling are linked to these dynamic changes of mitochondrial morphology.
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Liu, X., Kim, C. N., Yang, J., Jemmerson, R. & Wang, X. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86, 147–157 (1996).
Frank, S. et al. The role of dynamin-related protein 1, a mediator of mitochondrial fission, in apoptosis. Dev. Cell 1, 515–525 (2001).
Scorrano, L. et al. A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis. Dev. Cell 2, 55–67 (2002). References 2 and 3 provide the first demonstration that mitochondrial dynamics are essential for apoptosis.
Gomes, L. C., Di, B. G. & Scorrano, L. During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598 (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).
Khacho, M. et al. Mitochondrial dynamics impacts stem cell identity and fate decisions by regulating a nuclear transcriptional program. Cell Stem Cell 19, 232–247 (2016).
Kasahara, A., Cipolat, S., Chen, Y., Dorn, G. W. & Scorrano, L. Mitochondrial fusion directs cardiomyocyte differentiation via calcineurin and Notch signaling. Science 342, 734–737 (2013).
Yasukawa, K. et al. Mitofusin 2 inhibits mitochondrial antiviral signaling. Sci. Signal. 2, ra47 (2009).
Benda, C. Ueber die Spermatogenese der Vertebraten und höherer Evertebraten, II. Theil: die Histiogenese der Spermien. Arch. Anat. Physiol. 73, 393–398 (1898).
Collins, T. J., Berridge, M. J., Lipp, P. & Bootman, M. D. Mitochondria are morphologically and functionally heterogeneous within cells. EMBO J. 21, 1616–1627 (2002).
Kasahara, A. & Scorrano, L. Mitochondria: from cell death executioners to regulators of cell differentiation. Trends Cell Biol. 24, 761–770 (2014).
Pernas, L. & Scorrano, L. Mito-morphosis: mitochondrial fusion, fission, and cristae remodeling as key mediators of cellular function. Annu. Rev. Physiol. 78, 505–531 (2016).
Frey, T. G. & Mannella, C. A. The internal structure of mitochondria. Trends Biochem. Sci. 25, 319–324 (2000).
Vogel, F., Bornhovd, C., Neupert, W. & Reichert, A. S. Dynamic subcompartmentalization of the mitochondrial inner membrane. J. Cell Biol. 175, 237–247 (2006).
Gilkerson, R. W., Selker, J. M. & Capaldi, R. A. The cristal membrane of mitochondria is the principal site of oxidative phosphorylation. FEBS Lett. 546, 355–358 (2003).
Demongeot, J., Glade, N., Hansen, O. & Moreira, A. An open issue: the inner mitochondrial membrane (IMM) as a free boundary problem. Biochimie 89, 1049–1057 (2007).
Wolf, D. M. et al. Individual cristae within the same mitochondrion display different membrane potentials and are functionally independent. EMBO J. 38, e101056 (2019). Demonstration that individual cristae can be regarded as independent mitochondrial subcompartments.
Hackenbrock, C. R. Ultrastructural bases for metabolically linked mechanical activity in mitochondria. I. reversible ultrastructural changes with change in metabolic steady state in isolated liver mitochondria. J. Cell Biol. 30, 269–297 (1966). A classic article showing the link between mitochondrial morphological changes and function.
Mannella, C. A. et al. Topology of the mitochondrial inner membrane: dynamics and bioenergetic implications. IUBMB Life 52, 93–100 (2001).
Hackenbrock, C. R., Schneider, H., Lemasters, J. J. & Hochli, M. Relationships between bilayer lipid, motional freedom of oxidoreductase components, and electron transfer in the mitochondrial inner membrane. Adv. Exp. Med. Biol. 132, 245–263 (1980).
Cogliati, S. et al. Mitochondrial cristae shape determines respiratory chain supercomplexes assembly and respiratory efficiency. Cell 155, 160–171 (2013).
Chandel, N. S. Evolution of mitochondria as signaling organelles. Cell Metab. 22, 204–206 (2015).
Bahat, A. & Gross, A. Mitochondrial plasticity in cell fate regulation. J. Biol. Chem. 294, 13852–13863 (2019).
Nguyen, M., Millar, D. G., Yong, V. W., Korsmeyer, S. J. & Shore, G. C. Targeting of Bcl-2 to the mitochondrial outer membrane by a COOH-terminal signal anchor sequence. J. Biol. Chem. 268, 25265–25268 (1993).
Scorrano, L. & Korsmeyer, S. J. Mechanisms of cytochrome c release by proapoptotic BCL-2 family members. Biochem. Biophys. Res. Commun. 304, 437–444 (2003).
Karbowski, M., Norris, K. L., Cleland, M. M., Jeong, S. Y. & Youle, R. J. Role of Bax and Bak in mitochondrial morphogenesis. Nature 443, 658–662 (2006).
Karbowski, M. et al. Spatial and temporal association of Bax with mitochondrial fission sites, Drp1, and Mfn2 during apoptosis. J. Cell Biol. 159, 931–938 (2002).
Landes, T. et al. The BH3-only Bnip3 binds to the dynamin Opa1 to promote mitochondrial fragmentation and apoptosis by distinct mechanisms. EMBO Rep. 11, 459–465 (2010).
Lefkimmiatis, K., Leronni, D. & Hofer, A. M. The inner and outer compartments of mitochondria are sites of distinct cAMP/PKA signaling dynamics. J. Cell Biol. 202, 453–462 (2013).
Burdyga, A. et al. Phosphatases control PKA-dependent functional microdomains at the outer mitochondrial membrane. Proc. Natl Acad. Sci. USA 115, E6497–E6506 (2018).
Means, C. K. et al. An entirely specific type I A-kinase anchoring protein that can sequester two molecules of protein kinase A at mitochondria. Proc. Natl Acad. Sci. USA 108, E1227–E1235 (2011).
Castanier, C., Garcin, D., Vazquez, A. & Arnoult, D. Mitochondrial dynamics regulate the RIG-I-like receptor antiviral pathway. EMBO Rep. 11, 133–138 (2010).
Graham, T. R. & Kozlov, M. M. Interplay of proteins and lipids in generating membrane curvature. Curr. Opin. Cell Biol. 22, 430–436 (2010).
Friedman, J. R., Mourier, A., Yamada, J., McCaffery, J. M. & Nunnari, J. MICOS coordinates with respiratory complexes and lipids to establish mitochondrial inner membrane architecture. eLife 4, e07739 (2015).
Guarani, V. et al. QIL1 is a novel mitochondrial protein required for MICOS complex stability and cristae morphology. eLife 4, e06265 (2015).
Huynen, M. A., Muhlmeister, M., Gotthardt, K., Guerrero-Castillo, S. & Brandt, U. Evolution and structural organization of the mitochondrial contact site (MICOS) complex and the mitochondrial intermembrane space bridging (MIB) complex. Biochim. Biophys. Acta 1863, 91–101 (2015).
Barbot, M. et al. Mic10 oligomerizes to bend mitochondrial inner membranes at cristae junctions. Cell Metab. 21, 756–763 (2015).
Bohnert, M. et al. Central role of Mic10 in the mitochondrial contact site and cristae organizing system. Cell Metab. 21, 747–755 (2015).
Rabl, R. et al. Formation of cristae and crista junctions in mitochondria depends on antagonism between Fcj1 and Su e/g. J. Cell Biol. 185, 1047–1063 (2009). Identification of the role of MIC60 in cristae biogenesis.
Zerbes, R. M. et al. Role of MINOS in mitochondrial membrane architecture: cristae morphology and outer membrane interactions differentially depend on mitofilin domains. J. Mol. Biol. 422, 183–191 (2012).
John, G. B. et al. The mitochondrial inner membrane protein mitofilin controls cristae morphology. Mol. Biol. Cell 16, 1543–1554 (2005).
Hessenberger, M. et al. Regulated membrane remodeling by Mic60 controls formation of mitochondrial crista junctions. Nat. Commun. 8, 15258 (2017).
Tarasenko, D. et al. The MICOS component Mic60 displays a conserved membrane-bending activity that is necessary for normal cristae morphology. J. Cell Biol. 216, 889–899 (2017).
Eydt, K., Davies, K. M., Behrendt, C., Wittig, I. & Reichert, A. S. Cristae architecture is determined by an interplay of the MICOS complex and the F1FO ATP synthase via Mic27 and Mic10. Microb. Cell 4, 259–272 (2017).
Darshi, M. et al. ChChd3, an inner mitochondrial membrane protein, is essential for maintaining crista integrity and mitochondrial function. J. Biol. Chem. 286, 2918–2932 (2011).
Davies, K. M., Anselmi, C., Wittig, I., Faraldo-Gomez, J. D. & Kuhlbrandt, 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).
Strauss, M., Hofhaus, G., Schroder, R. R. & Kuhlbrandt, W. Dimer ribbons of ATP synthase shape the inner mitochondrial membrane. EMBO J. 27, 1154–1160 (2008).
Paumard, P. et al. The ATP synthase is involved in generating mitochondrial cristae morphology. EMBO J. 21, 221–230 (2002). First demonstration of a role for ATP synthase in mitochondrial cristae morphology.
Campanella, M. et al. Regulation of mitochondrial structure and function by the F1Fo-ATPase inhibitor protein, IF1. Cell Metab. 8, 13–25 (2008).
Bornhovd, C., Vogel, F., Neupert, W. & Reichert, A. S. Mitochondrial membrane potential is dependent on the oligomeric state of F1F0-ATP synthase supracomplexes. J. Biol. Chem. 281, 13990–13998 (2006).
Frezza, C. et al. OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion. Cell 126, 177–189 (2006).
Griparic, L., van der Wel, N. N., Orozco, I. J., Peters, P. J. & van der Bliek, A. M. Loss of the intermembrane space protein Mgm1/OPA1 induces swelling and localized constrictions along the lengths of mitochondria. J. Biol. Chem. 279, 18792–18798 (2004).
Glytsou, C. et al. Optic atrophy 1 is epistatic to the core MICOS component MIC60 in mitochondrial cristae shape control. Cell Rep. 17, 3024–3034 (2016).
Quintana-Cabrera, R. et al. The cristae modulator optic atrophy 1 requires mitochondrial ATP synthase oligomers to safeguard mitochondrial function. Nat. Commun. 9, 3399 (2018).
Faelber, K. et al. Structure and assembly of the mitochondrial membrane remodelling GTPase Mgm1. Nature 571, 429–433 (2019).
Meeusen, S. et al. Mitochondrial inner-membrane fusion and crista maintenance requires the dynamin-related GTPase Mgm1. Cell 127, 383–395 (2006).
Amutha, B., Gordon, D. M., Gu, Y. & Pain, D. A novel role of Mgm1p, a dynamin-related GTPase, in ATP synthase assembly and cristae formation/maintenance. Biochem. J. 381, 19–23 (2004).
Sesaki, H., Southard, S. M., Yaffe, M. P. & Jensen, R. E. Mgm1p, a dynamin-related GTPase, is essential for fusion of the mitochondrial outer membrane. Mol. Biol. Cell 14, 2342–2356 (2003).
Wang, L. et al. FAM92A1 is a BAR domain protein required for mitochondrial ultrastructure and function. J. Cell Biol. 218, 97–111 (2019).
Chen, H. et al. Titration of mitochondrial fusion rescues Mff-deficient cardiomyopathy. J. Cell Biol. 211, 795–805 (2015).
Labbe, K., Murley, A. & Nunnari, J. Determinants and functions of mitochondrial behavior. Annu. Rev. Cell Dev. Biol. 30, 357–391 (2014).
Friedman, J. R. & Nunnari, J. Mitochondrial form and function. Nature 505, 335–343 (2014).
Tilokani, L., Nagashima, S., Paupe, V. & Prudent, J. Mitochondrial dynamics: overview of molecular mechanisms. Essays Biochem. 62, 341–360 (2018).
Zuchner, S. et al. Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat. Genet. 36, 449–451 (2004).
Santel, A. & Fuller, M. T. Control of mitochondrial morphology by a human mitofusin. J. Cell Sci. 114, 867–874 (2001).
Alexander, C. et al. OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat. Genet. 26, 211–215 (2000).
Delettre, C. et al. Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat. Genet. 26, 207–210 (2000). References 66 and 67 provide the first demonstration of a genetic disorder caused by mutant mitochondria-shaping proteins.
Cipolat, S. O., Martins de Brito, O., Dal Zilio, B. & Scorrano, L. OPA1 requires mitofusin 1 to promote mitochondrial fusion. Proc. Natl Acad. Sci. USA 101, 15927–15932 (2004).
Rojo, M., Legros, F., Chateau, D. & Lombes, A. Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo. J. Cell Sci. 115, 1663–1674 (2002).
Ishihara, N., Eura, Y. & Mihara, K. Mitofusin 1 and 2 play distinct roles in mitochondrial fusion reactions via GTPase activity. J. Cell Sci. 117, 6535–6546 (2004).
de Brito, O. M. & Scorrano, L. Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature 456, 605–610 (2008). Identification of the first structural ER–mitochondria tether in mammals.
Qi, Y. et al. Structures of human mitofusin 1 provide insight into mitochondrial tethering. J. Cell Biol. 215, 621–629 (2016).
Brandt, T., Cavellini, L., Kühlbrandt, W. & Cohen, M. M. A mitofusin-dependent docking ring complex triggers mitochondrial fusion in vitro. eLife 5, e14618 (2016).
Ban, T. et al. Molecular basis of selective mitochondrial fusion by heterotypic action between OPA1 and cardiolipin. Nat. Cell Biol. 19, 856–863 (2017).
Herlan, M., Vogel, F., Bornhovd, C., Neupert, W. & Reichert, A. S. Processing of Mgm1 by the rhomboid-type protease Pcp1 is required for maintenance of mitochondrial morphology and of mitochondrial DNA. J. Biol. Chem. 278, 27781–27788 (2003).
McQuibban, G. A., Saurya, S. & Freeman, M. Mitochondrial membrane remodelling regulated by a conserved rhomboid protease. Nature 423, 537–541 (2003).
DeVay, R. M. et al. Coassembly of Mgm1 isoforms requires cardiolipin and mediates mitochondrial inner membrane fusion. J. Cell Biol. 186, 793–803 (2009).
Griparic, L., Kanazawa, T. & van der Bliek, A. M. Regulation of the mitochondrial dynamin-like protein Opa1 by proteolytic cleavage. J. Cell Biol. 178, 757–764 (2007).
Baker, M. J. et al. Stress-induced OMA1 activation and autocatalytic turnover regulate OPA1-dependent mitochondrial dynamics. EMBO J. 33, 578–593 (2014).
Ehses, S. et al. Regulation of OPA1 processing and mitochondrial fusion by m-AAA protease isoenzymes and OMA1. J. Cell Biol. 187, 1023–1036 (2009).
Head, B., Griparic, L., Amiri, M., Gandre-Babbe, S. & van der Bliek, A. M. Inducible proteolytic inactivation of OPA1 mediated by the OMA1 protease in mammalian cells. J. Cell Biol. 187, 959–966 (2009).
Cipolat, S. et al. Mitochondrial rhomboid PARL regulates cytochrome c release during apoptosis via OPA1-dependent cristae remodeling. Cell 126, 163–175 (2006).
Saita, S. et al. PARL mediates Smac proteolytic maturation in mitochondria to promote apoptosis. Nat. Cell Biol. 19, 318–328 (2017).
Botham, A. et al. Global interactome mapping of mitochondrial intermembrane space proteases identifies a novel function for HTRA2. Proteomics 19, 1900139 (2019).
Smirnova, E., Griparic, L., Shurland, D. L. & van der Bliek, A. M. Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells. Mol. Biol. Cell 12, 2245–2256 (2001).
Otera, H. et al. Mff is an essential factor for mitochondrial recruitment of Drp1 during mitochondrial fission in mammalian cells. J. Cell Biol. 191, 1141–1158 (2010).
James, D. I., Parone, P. A., Mattenberger, Y. & Martinou, J. C. hFis1, a novel component of the mammalian mitochondrial fission machinery. J. Biol. Chem. 278, 36373–36379 (2003).
Palmer, C. S. et al. MiD49 and MiD51, new components of the mitochondrial fission machinery. EMBO Rep. 12, 565–573 (2011).
Loson, O. C., Song, Z., Chen, H. & Chan, D. C. Fis1, Mff, MiD49, and MiD51 mediate Drp1 recruitment in mitochondrial fission. Mol. Biol. Cell 24, 659–667 (2013).
Kalia, R. et al. Structural basis of mitochondrial receptor binding and constriction by DRP1. Nature 558, 401–405 (2018).
Griffin, E. E., Graumann, J. & Chan, D. C. The WD40 protein Caf4p is a component of the mitochondrial fission machinery and recruits Dnm1p to mitochondria. J. Cell Biol. 170, 237–248 (2005).
Karren, M. A., Coonrod, E. M., Anderson, T. K. & Shaw, J. M. The role of Fis1p-Mdv1p interactions in mitochondrial fission complex assembly. J. Cell Biol. 171, 291–301 (2005).
Naylor, K. et al. MDV1 interacts with assembled DNM1 to promote mitochondrial division. J. Biol. Chem. 281, 2177–2183 (2006).
Alirol, E. et al. The mitochondrial fission protein hFis1 requires the endoplasmic reticulum gateway to induce apoptosis. Mol. Biol. Cell 17, 4593–4605 (2006).
Wong, Y. C., Ysselstein, D. & Krainc, D. Mitochondria–lysosome contacts regulate mitochondrial fission via RAB7 GTP hydrolysis. Nature 554, 382 (2018).
Yu, R., Jin, S.-B., Lendahl, U., Nistér, M. & Zhao, J. Human Fis1 regulates mitochondrial dynamics through inhibition of the fusion machinery. EMBO J. 38, e99748 (2019).
Ishihara, N. et al. Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nat. Cell Biol. 11, 958–966 (2009).
Osellame, L. D. et al. Cooperative and independent roles of the Drp1 adaptors Mff, MiD49 and MiD51 in mitochondrial fission. J. Cell Sci. 129, 2170–2181 (2016).
Bohuszewicz, O. & Low, H. H. Structure of a mitochondrial fission dynamin in the closed conformation. Nat. Struct. Mol. Biol. 25, 722–731 (2018).
Lee, J. E., Westrate, L. M., Wu, H., Page, C. & Voeltz, G. K. Multiple dynamin family members collaborate to drive mitochondrial division. Nature 540, 139–143 (2016).
Fonseca, T. B., Sánchez-Guerrero, Á., Milosevic, I. & Raimundo, N. Mitochondrial fission requires DRP1 but not dynamins. Nature 570, E34–E42 (2019).
Kamerkar, S. C., Kraus, F., Sharpe, A. J., Pucadyil, T. J. & Ryan, M. T. Dynamin-related protein 1 has membrane constricting and severing abilities sufficient for mitochondrial and peroxisomal fission. Nat. Commun. 9, 5239 (2018).
Cribbs, J. T. & Strack, S. Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein kinase and calcineurin regulates mitochondrial fission and cell death. EMBO Rep. 8, 939–944 (2007).
Chang, C. R. & Blackstone, C. Cyclic AMP-dependent protein kinase phosphorylation of Drp1 regulates its GTPase activity and mitochondrial morphology. J. Biol. Chem. 282, 21583–21587 (2007). References 103 and 104 provide the first evidence for a functional PTM in a mitochondria-shaping protein.
Cereghetti, G. M. et al. Dephosphorylation by calcineurin regulates translocation of Drp1 to mitochondria. Proc. Natl Acad. Sci. USA 105, 15803–15808 (2008).
Yu, R. et al. The phosphorylation status of Ser-637 in dynamin-related protein 1 (Drp1) does not determine Drp1 recruitment to mitochondria. J. Biol. Chem. 294, 17262–17277 (2019).
Friedman, J. R. et al. ER tubules mark sites of mitochondrial division. Science 334, 358–362 (2011). Identification of the association between ER and mitochondrial fission sites.
Iwasawa, R., Mahul-Mellier, A. L., Datler, C., Pazarentzos, E. & Grimm, S. Fis1 and Bap31 bridge the mitochondria-ER interface to establish a platform for apoptosis induction. EMBO J. 30, 556–568 (2011).
Rusinol, A. E., Cui, Z., Chen, M. H. & Vance, J. E. A unique mitochondria-associated membrane fraction from rat liver has a high capacity for lipid synthesis and contains pre-Golgi secretory proteins including nascent lipoproteins. J. Biol. Chem. 269, 27494–27502 (1994). Discovery of mitochondria-associated ER membranes.
Giacomello, M. & Pellegrini, L. The coming of age of the mitochondria-ER contact: a matter of thickness. Cell Death Differ. 23, 1417–1427 (2016).
De Mario, A., Quintana-Cabrera, R., Martinvalet, D. & Giacomello, M. (Neuro)degenerated mitochondria-ER contacts. Biochem. Biophys. Res. Commun. 483, 1096–1109 (2017).
Friedman, J. R., Webster, B. M., Mastronarde, D. N., Verhey, K. J. & Voeltz, G. K. ER sliding dynamics and ER-mitochondrial contacts occur on acetylated microtubules. J. Cell Biol. 190, 363–375 (2010).
Korobova, F., Ramabhadran, V. & Higgs, H. N. An actin-dependent step in mitochondrial fission mediated by the ER-associated formin INF2. Science 339, 464–467 (2013).
Manor, U. et al. A mitochondria-anchored isoform of the actin-nucleating spire protein regulates mitochondrial division. eLife 4, e08828 (2015).
Chakrabarti, R. et al. INF2-mediated actin polymerization at the ER stimulates mitochondrial calcium uptake, inner membrane constriction, and division. J. Cell Biol. 217, 251–268 (2017).
Wong, L. H., Gatta, A. T. & Levine, T. P. Lipid transfer proteins: the lipid commute via shuttles, bridges and tubes. Nat. Rev. Mol. Cell Biol. 20, 85–101 (2019).
Janer, A. et al. SLC25A46 is required for mitochondrial lipid homeostasis and cristae maintenance and is responsible for Leigh syndrome. EMBO Mol. Med. 8, 1019–1038 (2016).
Wu, M. J. et al. Epithelial-mesenchymal transition directs stem cell polarity via regulation of mitofusin. Cell Metab. 29, 993–1002.e6 (2019).
Singaravelu, K. et al. Mitofusin 2 Regulates STIM1 migration from the Ca2+ store to the plasma membrane in cells with depolarized mitochondria. J. Biol. Chem. 286, 12189–12201 (2011).
Ping, H. A., Kraft, L. M., Chen, W., Nilles, A. E. & Lackner, L. L. Num1 anchors mitochondria to the plasma membrane via two domains with different lipid binding specificities. J. Cell Biol. 213, 513–524 (2016).
Steinman, R. M., Mellman, I. S., Muller, W. A. & Cohn, Z. A. Endocytosis and the recycling of plasma membrane. J. Cell Biol. 96, 1–27 (1983).
Huotari, J. & Helenius, A. Endosome maturation. EMBO J. 30, 3481–3500 (2011).
Das, A., Nag, S., Mason, A. B. & Barroso, M. M. Endosome-mitochondria interactions are modulated by iron release from transferrin. J. Cell Biol. 214, 831–845 (2016).
Charman, M., Kennedy, B. E., Osborne, N. & Karten, B. MLN64 mediates egress of cholesterol from endosomes to mitochondria in the absence of functional Niemann-Pick type C1 protein. J. Lipid Res. 51, 1023–1034 (2010).
Hsu, F. et al. Rab5 and Alsin regulate stress-activated cytoprotective signaling on mitochondria. eLife 7, e32282 (2018).
Wang, H. et al. Perilipin 5, a lipid droplet-associated protein, provides physical and metabolic linkage to mitochondria. J. Lipid Res. 52, 2159–2168 (2011).
Benador, I. Y. et al. Mitochondria bound to lipid droplets have unique bioenergetics, composition, and dynamics that support lipid droplet expansion. Cell Metab. 27, 869–885.e6 (2018).
Hammerschmidt, P. et al. CerS6-derived sphingolipids interact with Mff and promote mitochondrial fragmentation in obesity. Cell 177, 1536–1552.e23 (2019).
Fransen, M., Lismont, C. & Walton, P. The peroxisome-mitochondria connection: how and why? Int. J. Mol. Sci. 18, E1126 (2017).
Shai, N. et al. Systematic mapping of contact sites reveals tethers and a function for the peroxisome-mitochondria contact. Nat. Commun. 9, 1761 (2018).
Daniele, T. et al. Mitochondria and melanosomes establish physical contacts modulated by Mfn2 and involved in organelle biogenesis. Curr. Biol. 24, 393–403 (2014).
Mattiazzi Ušaj, M. et al. Genome-wide localization study of yeast Pex11 identifies peroxisome–mitochondria interactions through the ERMES complex. J. Mol. Biol. 427, 2072–2087 (2015).
McGuinness, M. C. et al. Role of ALDP (ABCD1) and mitochondria in X-linked adrenoleukodystrophy. Mol. Cell. Biol. 23, 744–753 (2003).
Fan, J., Li, X., Issop, L., Culty, M. & Papadopoulos, V. ACBD2/ECI2-mediated peroxisome-mitochondria interactions in Leydig cell steroid biosynthesis. Mol. Endocrinol. 30, 763–782 (2016).
Kerr, J. F., Wyllie, A. H. & Currie, A. R. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br. J. Cancer. 26, 239–257 (1972).
Karbowski, M. et al. Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J. Cell Biol. 164, 493–499 (2004).
Wasiak, S., Zunino, R. & McBride, H. M. Bax/Bak promote sumoylation of DRP1 and its stable association with mitochondria during apoptotic cell death. J. Cell Biol. 177, 439–450 (2007).
Braschi, E., Zunino, R. & McBride, H. M. MAPL is a new mitochondrial SUMO E3 ligase that regulates mitochondrial fission. EMBO Rep. 10, 748–754 (2009).
Figueroa-Romero, C. et al. SUMOylation of the mitochondrial fission protein Drp1 occurs at multiple nonconsensus sites within the B domain and is linked to its activity cycle. FASEB J. 23, 3917–3927 (2009).
Prudent, J. et al. MAPL SUMOylation of Drp1 stabilizes an ER/mitochondrial platform required for cell death. Mol. Cell 59, 941–955 (2015).
Guo, X. et al. Inhibition of mitochondrial fragmentation diminishes Huntington’s disease-associated neurodegeneration. J. Clin. Invest. 123, 5371–5388 (2013).
Qi, X., Qvit, N., Su, Y. C. & Mochly-Rosen, D. A novel Drp1 inhibitor diminishes aberrant mitochondrial fission and neurotoxicity. J. Cell Sci. 126, 789–802 (2013).
Merrill, R. A., Slupe, A. M. & Strack, S. N-terminal phosphorylation of protein phosphatase-2A/Bβ2 regulates translocation to mitochondria, dynamin-related protein 1 dephosphorylation, and neuronal survival. FEBS J. 280, 662–673 (2013).
Costa, V. et al. Mitochondrial fission and cristae disruption increase the response of cell models of Huntington’s disease to apoptotic stimuli. EMBO Mol. Med. 2, 490–503 (2010).
Cho, D. H. et al. S-nitrosylation of Drp1 mediates beta-amyloid-related mitochondrial fission and neuronal injury. Science 324, 102–105 (2009).
Liot, G. et al. Complex II inhibition by 3-NP causes mitochondrial fragmentation and neuronal cell death. Cell Death Differ. 16, 899–909 (2009).
Brooks, C., Wei, Q., Cho, S. G. & Dong, Z. Regulation of mitochondrial dynamics in acute kidney injury in cell culture and rodent models. J. Clin. Invest 119, 1275–1285 (2009).
Zhang, Z., Liu, L., Wu, S. & Xing, D. Drp1, Mff, Fis1, and MiD51 are coordinated to mediate mitochondrial fission during UV irradiation–induced apoptosis. FASEB J. 30, 466–476 (2016).
Li, G. et al. Mitochondrial translocation and interaction of cofilin and Drp1 are required for erucin-induced mitochondrial fission and apoptosis. Oncotarget 6, 1834–1849 (2015).
Chou, C. H. et al. GSK3beta-mediated Drp1 phosphorylation induced elongated mitochondrial morphology against oxidative stress. PLoS One 7, e49112 (2012).
Torres, G. et al. Glucagon-like peptide-1 inhibits vascular smooth muscle cell dedifferentiation through mitochondrial dynamics regulation. Biochem. Pharmacol. 104, 52–61 (2016).
Li, A. et al. Metformin and resveratrol inhibit Drp1-mediated mitochondrial fission and prevent ER stress-associated NLRP3 inflammasome activation in the adipose tissue of diabetic mice. Mol. Cell. Endocrinol. 434, 36–47 (2016).
Lavie, J. et al. Mitochondrial morphology and cellular distribution are altered in SPG31 patients and are linked to DRP1 hyperphosphorylation. Hum. Mol. Genet. 26, 674–685 (2016).
Li, J. et al. Pharmacological activation of AMPK prevents Drp1-mediated mitochondrial fission and alleviates endoplasmic reticulum stress-associated endothelial dysfunction. J. Mol. Cell. Cardiol. 86, 62–74 (2015).
Sharp, W. W. et al. Dynamin-related protein 1 (Drp1)-mediated diastolic dysfunction in myocardial ischemia-reperfusion injury: therapeutic benefits of Drp1 inhibition to reduce mitochondrial fission. FASEB J. 28, 316–326 (2014).
Kim, B. et al. Inhibition of Drp1-dependent mitochondrial division impairs myogenic differentiation. Am. J. Physiol. Regul. Integr. Comp. Physiol. 305, R927–R938 (2013).
Cereghetti, G. M., Costa, V. & Scorrano, L. Inhibition of Drp1-dependent mitochondrial fragmentation and apoptosis by a polypeptide antagonist of calcineurin. Cell Death Differ. 17, 1785–1794 (2010).
Ferreira, J. C. B. et al. A selective inhibitor of mitofusin 1-βIIPKC association improves heart failure outcome in rats. Nat. Commun. 10, 329 (2019).
Leboucher, Guillaume P. et al. Stress-induced phosphorylation and proteasomal degradation of mitofusin 2 facilitates mitochondrial fragmentation and apoptosis. Mol. Cell 47, 547–557 (2012).
Shutt, T., Geoffrion, M., Milne, R. & McBride, H. M. The intracellular redox state is a core determinant of mitochondrial fusion. EMBO Rep. 13, 909–915 (2012).
Mattie, S., Riemer, J., Wideman, J. G. & McBride, H. M. A new mitofusin topology places the redox-regulated C terminus in the mitochondrial intermembrane space. J. Cell Biol. 217, 507–515 (2018). This identification of a revised topology for mitofusin with the coiled coil 2 region exposed to the intermembrane space has had profound implications for the mechanism of mitochondrial fusion and the development of mitofusin-targeting drugs.
Cogliati, S., Enriquez, J. A. & Scorrano, L. Mitochondrial cristae: where beauty meets functionality. Trends Biochem. Sci. 41, 261–273 (2016).
Yamaguchi, R. et al. Opa1-mediated cristae opening is Bax/Bak and BH3 dependent, required for apoptosis, and independent of Bak oligomerization. Mol. Cell 31, 557–569 (2008).
Chen, X. et al. Targeting mitochondrial structure sensitizes acute myeloid leukemia to venetoclax treatment. Cancer Discov. 9, 890–909 (2019).
Wu, W. et al. OPA1 overexpression ameliorates mitochondrial cristae remodeling, mitochondrial dysfunction, and neuronal apoptosis in prion diseases. Cell Death Dis. 10, 710–710 (2019).
Varanita, T. et al. The OPA1-dependent mitochondrial cristae remodeling pathway controls atrophic, apoptotic, and ischemic tissue damage. Cell Metab. 21, 834–844 (2015).
Ramonet, D. et al. Optic atrophy 1 mediates mitochondria remodeling and dopaminergic neurodegeneration linked to complex I deficiency. Cell Death Differ. 20, 77–85 (2012).
Faccenda, D. et al. Control of mitochondrial remodeling by the ATPase inhibitory factor 1 unveils a pro-survival relay via OPA1. Cell Rep. 18, 1869–1883 (2017).
Sun, M. G. et al. Correlated three-dimensional light and electron microscopy reveals transformation of mitochondria during apoptosis. Nat. Cell Biol. 9, 1057–1065 (2007).
Otera, H., Miyata, N., Kuge, O. & Mihara, K. Drp1-dependent mitochondrial fission via MiD49/51 is essential for apoptotic cristae remodeling. J. Cell Biol. 212, 531–544 (2016).
Germain, M., Mathai, J. P., McBride, H. M. & Shore, G. C. Endoplasmic reticulum BIK initiates DRP1-regulated remodelling of mitochondrial cristae during apoptosis. EMBO J. 24, 1546–1556 (2005).
Bernardi, P., Rasola, A., Forte, M. & Lippe, G. The mitochondrial permeability transition pore: channel formation by F-ATP synthase, integration in signal transduction, and role in pathophysiology. Physiol. Rev. 95, 1111–1155 (2015).
Samant, S. A. et al. SIRT3 deacetylates and activates OPA1 to regulate mitochondrial dynamics during stress. Mol. Cell. Biol. 34, 807–819 (2014).
Bossy, B. et al. S-Nitrosylation of DRP1 does not affect enzymatic activity and is not specific to Alzheimer’s disease. J. Alzheimers. Dis. 20, S513–S526 (2010).
Makino, A. et al. Regulation of mitochondrial morphology and function by O-GlcNAcylation in neonatal cardiac myocytes. Am. J. Physiol. Regul. Integr. Comp. Physiol. 300, R1296–R1302 (2011).
Pickles, S., Vigié, P. & Youle, R. J. Mitophagy and quality control mechanisms in mitochondrial maintenance. Curr. Biol. 28, R170–R185 (2018).
Mottis, A., Jovaisaite, V. & Auwerx, J. The mitochondrial unfolded protein response in mammalian physiology. Mamm. Genome 25, 424–433 (2014).
Anand, R., Langer, T. & Baker, M. J. Proteolytic control of mitochondrial function and morphogenesis. Biochim. Biophys. Acta 1833, 195–204 (2013).
Sugiura, A., McLelland, G. L., Fon, E. A. & McBride, H. M. A new pathway for mitochondrial quality control: mitochondrial-derived vesicles. EMBO J. 33, 2142–2156 (2014). Discovery and characterization of the role of MDVs in quality control of mitochondria.
Eisner, V., Picard, M. & Hajnóczky, G. Mitochondrial dynamics in adaptive and maladaptive cellular stress responses. Nat. Cell Biol. 20, 755–765 (2018).
Ziviani, E., Tao, R. N. & Whitworth, A. J. Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc. Natl Acad. Sci. USA 107, 5018–5023 (2010). This article demonstrates that parkin ubiquitylates the mitochondrial fusion protein mitofusin, providing a first mechanistic link between mitophagy and mitochondrial morphology.
Kitada, T. et al. Mutations in the parkin gene cause autosomal recessive juvenile parkinsonism. Nature 392, 605–608 (1998).
Narendra, D., Tanaka, A., Suen, D. F. & Youle, R. J. Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. J. Cell Biol. 183, 795–803 (2008). This article demonstrates that parkin translocates to dysfunctional mitochondria, providing the first mechanistic explanation for selective autophagy.
Park, J., Lee, G. & Chung, J. The PINK1-parkin pathway is involved in the regulation of mitochondrial remodeling process. Biochem. Biophys. Res. Commun. 378, 518–523 (2009).
Sarraf, S. A. et al. Landscape of the PARKIN-dependent ubiquitylome in response to mitochondrial depolarization. Nature 496, 372 (2013).
Sugiura, A. et al. MITOL regulates endoplasmic reticulum-mitochondria contacts via mitofusin2. Mol. Cell 51, 20–34 (2013).
Chen, Y. & Dorn, G. W. II PINK1-phosphorylated mitofusin 2 is a parkin receptor for culling damaged mitochondria. Science 340, 471–475 (2013).
Toyama, E. Q. et al. AMP-activated protein kinase mediates mitochondrial fission in response to energy stress. Science 351, 275–281 (2016).
Niemann, A., Ruegg, M., La Padula, V., Schenone, A. & Suter, U. Ganglioside-induced differentiation associated protein 1 is a regulator of the mitochondrial network: new implications for Charcot-Marie-Tooth disease. J. Cell Biol. 170, 1067–1078 (2005).
Neuspiel, M. et al. Cargo-selected transport from the mitochondria to peroxisomes is mediated by vesicular carriers. Curr. Biol. 18, 102–108 (2008).
Hughes, A. L., Hughes, C. E., Henderson, K. A., Yazvenko, N. & Gottschling, D. E. Selective sorting and destruction of mitochondrial membrane proteins in aged yeast. eLife 5, e13943 (2016).
Mitchell, P. & Moyle, J. Chemiosmotic hypothesis of oxidative phosphorylation. Nature 213, 137–139 (1967).
Patten, D. A. et al. OPA1-dependent cristae modulation is essential for cellular adaptation to metabolic demand. EMBO J. 33, 2676–2691 (2014).
Miyamoto, T. et al. Compartmentalized AMPK signaling illuminated by genetically encoded molecular sensors and actuators. Cell Rep. 11, 657–670 (2015).
Chouchani, E. T. et al. A unifying mechanism for mitochondrial superoxide production during ischemia-reperfusion injury. Cell Metab. 23, 254–263 (2016).
Zhou, H. et al. NR4A1 aggravates the cardiac microvascular ischemia reperfusion injury through suppressing FUNDC1-mediated mitophagy and promoting Mff-required mitochondrial fission by CK2α. Basic Res. Cardiol. 113, 23 (2018).
Mitra, K., Wunder, C., Roysam, B., Lin, G. & Lippincott-Schwartz, J. A hyperfused mitochondrial state achieved at G1-S regulates cyclin E buildup and entry into S phase. Proc. Natl Acad. Sci. USA 106, 11960–11965 (2009).
Taguchi, N., Ishihara, N., Jofuku, A., Oka, T. & Mihara, K. Mitotic phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial fission. J. Biol. Chem. 282, 11521–11529 (2007). This article identifies mitochondrial fission mediated by DRP1 as a crucial step in mitosis.
Chung, J. Y., Steen, J. A. & Schwarz, T. L. Phosphorylation-induced motor shedding is required at mitosis for proper distribution and passive inheritance of mitochondria. Cell Rep. 16, 2142–2155 (2016).
Kaplon, J., van Dam, L. & Peeper, D. Two-way communication between the metabolic and cell cycle machineries: the molecular basis. Cell Cycle 14, 2022–2032 (2015).
Kashatus, D. F. et al. RALA and RALBP1 regulate mitochondrial fission at mitosis. Nat. Cell Biol. 13, 1108 (2011).
Montemurro, C. et al. Cell cycle-related metabolism and mitochondrial dynamics in a replication-competent pancreatic beta-cell line. Cell Cycle 16, 2086–2099 (2017).
Wang, H. et al. Parkin ubiquitinates Drp1 for proteasome-dependent degradation: implication of dysregulated mitochondrial dynamics in Parkinson disease. J. Biol. Chem. 286, 11649–11658 (2011).
Horn, S. R. et al. Regulation of mitochondrial morphology by APC/CCdh1-mediated control of Drp1 stability. Mol. Biol. Cell 22, 1207–1216 (2011).
Zunino, R., Schauss, A., Rippstein, P., Andrade-Navarro, M. & McBride, H. M. The SUMO protease SENP5 is required to maintain mitochondrial morphology and function. J. Cell Sci. 120, 1178–1188 (2007).
Zunino, R., Braschi, E., Xu, L. & McBride, H. M. Translocation of SenP5 from the nucleoli to the mitochondria modulates DRP1-dependent fission during mitosis. J. Biol. Chem. 284, 17783–17795 (2009).
Zhong, X. et al. Mitochondrial dynamics is critical for the full pluripotency and embryonic developmental potential of pluripotent stem cells. Cell Metab. 29, 979–992.e4 (2019).
Luchsinger, L. L., de Almeida, M. J., Corrigan, D. J., Mumau, M. & Snoeck, H. W. Mitofusin 2 maintains haematopoietic stem cells with extensive lymphoid potential. Nature 529, 528–531 (2016).
Jung, S. et al. Mitofusin 2, a mitochondria-ER tethering protein, facilitates osteoclastogenesis by regulating the calcium-calcineurin-NFATc1 axis. Biochem. Biophys. Res. Commun. 516, 202–208 (2019).
Bahat, A. et al. MTCH2-mediated mitochondrial fusion drives exit from naïve pluripotency in embryonic stem cells. Nat. Commun. 9, 5132 (2018).
Favaro, G. et al. DRP1-mediated mitochondrial shape controls calcium homeostasis and muscle mass. Nat. Commun. 10, 2576 (2019).
Szabadkai, G. et al. Drp-1-dependent division of the mitochondrial network blocks intraorganellar Ca2+ waves and protects against Ca2+-mediated apoptosis. Mol. Cell 16, 59–68 (2004).
Herranz, N. & Gil, J. Mitochondria and senescence: new actors for an old play. EMBO J. 35, 701–702 (2016).
Chapman, J., Fielder, E. & Passos, J. F. Mitochondrial dysfunction and cell senescence: deciphering a complex relationship. FEBS Lett. 593, 1566–1579 (2019).
Vasileiou, P. V. S. et al. Mitochondrial homeostasis and cellular senescence. Cells 8, E686 (2019).
Tezze, C. et al. Age-associated loss of OPA1 in muscle impacts muscle mass, metabolic homeostasis, systemic inflammation, and epithelial senescence. Cell Metab. 25, 1374–1389 (2017).
Pereira, R. O. et al. OPA1 deficiency promotes secretion of FGF21 from muscle that prevents obesity and insulin resistance. EMBO J. 36, 2126–2145 (2017).
Rodriguez-Nuevo, A. et al. Mitochondrial DNA and TLR9 drive muscle inflammation upon Opa1 deficiency. EMBO J. 37, e96553 (2018).
Restelli, L. M. et al. Neuronal mitochondrial dysfunction activates the integrated stress response to induce fibroblast growth factor 21. Cell Rep. 24, 1407–1414 (2018).
Dogan, S. A. et al. Tissue-specific loss of DARS2 activates stress responses independently of respiratory chain deficiency in the heart. Cell Metab. 19, 458–469 (2014).
Schneeberger, M. et al. Mitofusin 2 in POMC neurons connects ER stress with leptin resistance and energy imbalance. Cell 155, 172–187 (2013).
Mancini, G. et al. Mitofusin 2 in mature adipocytes controls adiposity and body weight. Cell Rep. 26, 2849–2858.e2844 (2019).
Chung, K.-P. et al. Mitofusins regulate lipid metabolism to mediate the development of lung fibrosis. Nat. Commun. 10, 3390 (2019).
Franco, A. et al. Correcting mitochondrial fusion by manipulating mitofusin conformations. Nature 540, 74–79 (2016). Identification of the first pharmacological mitochondrial fusion activators.
Cassidy-Stone, A. et al. Chemical inhibition of the mitochondrial division dynamin reveals its role in Bax/Bak-dependent mitochondrial outer membrane permeabilization. Dev. Cell 14, 193–204 (2008).
Mallat, A. et al. Discovery and characterization of selective small molecule inhibitors of the mammalian mitochondrial division dynamin, DRP1. Biochem. Biophys. Res. Commun. 499, 556–562 (2018).
Quirós, P. M., Mottis, A. & Auwerx, J. Mitonuclear communication in homeostasis and stress. Nat. Rev. Mol. Cell Biol. 17, 213–226 (2016).
Anderson, G. R. et al. Dysregulation of mitochondrial dynamics proteins are a targetable feature of human tumors. Nat. Commun. 9, 1677 (2018).
Antonny, B. et al. Membrane fission by dynamin: what we know and what we need to know. EMBO J. 35, 2270–2284 (2016).
Chappie, J. S. et al. An intramolecular signaling element that modulates dynamin function in vitro and in vivo. Mol. Biol. Cell 20, 3561–3571 (2009).
Daumke, O., Roux, A. & Haucke, V. BAR domain scaffolds in dynamin-mediated membrane fission. Cell 156, 882–892 (2014).
Sochacki, K. A. & Taraska, J. W. From flat to curved clathrin: controlling a plastic ratchet. Trends Cell Biol. 29, 241–256 (2019).
Ford, M. G., Jenni, S. & Nunnari, J. The crystal structure of dynamin. Nature 477, 561–566 (2011).
Roux, A., Uyhazi, K., Frost, A. & De Camilli, P. GTP-dependent twisting of dynamin implicates constriction and tension in membrane fission. Nature 441, 528–531 (2006).
Rismanchi, N., Soderblom, C., Stadler, J., Zhu, P. P. & Blackstone, C. Atlastin GTPases are required for Golgi apparatus and ER morphogenesis. Hum. Mol. Genet. 17, 1591–1604 (2008).
Smith, J. J. & Aitchison, J. D. Peroxisomes take shape. Nat. Rev. Mol. Cell Biol. 14, 803–817 (2013).
McNew, J. A., Sondermann, H., Lee, T., Stern, M. & Brandizzi, F. GTP-dependent membrane fusion. Annu. Rev. Cell Dev. Biol. 29, 529–550 (2013).
Zuchner, S. et al. Mutations in the pleckstrin homology domain of dynamin 2 cause dominant intermediate Charcot-Marie-Tooth disease. Nat. Genet. 37, 289–294 (2005).
Rizzuto, R. et al. Close contacts with the endoplasmic reticulum as determinants of mitochondrial Ca2+ responses. Science 280, 1763–1766 (1998).
Giacomello, M. et al. Ca2+ hot spots on the mitochondrial surface are generated by Ca2+ mobilization from stores, but not by activation of store-operated Ca2+ channels. Mol. Cell 38, 280–290 (2010).
Csordas, G. et al. Imaging interorganelle contacts and local calcium dynamics at the ER-mitochondrial interface. Mol. Cell 39, 121–132 (2010).
Rizzuto, R., De, S. D., Raffaello, A. & Mammucari, C. Mitochondria as sensors and regulators of calcium signalling. Nat. Rev. Mol. Cell Biol. 13, 566–578 (2012).
Wu, S. et al. Binding of FUN14 domain containing 1 with inositol 1,4,5-trisphosphate receptor in mitochondria-associated endoplasmic reticulum membranes maintains mitochondrial dynamics and function in hearts in vivo. Circulation 136, 2248–2266 (2017).
Park, S. J. et al. DISC1 modulates neuronal stress responses by gate-keeping er-mitochondria Ca2+ transfer through the MAM. Cell Rep. 21, 2748–2759 (2017).
Simmen, T. et al. PACS-2 controls endoplasmic reticulum–mitochondria communication and Bid-mediated apoptosis. EMBO J. 24, 717–729 (2005).
Vance, J. E. Newly made phosphatidylserine and phosphatidylethanolamine are preferentially translocated between rat liver mitochondria and endoplasmic reticulum. J. Biol. Chem. 266, 89–97 (1991).
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).
Chen, Y. et al. Mitofusin 2-containing mitochondrial-reticular microdomains direct rapid cardiomyocyte bioenergetic responses via interorganelle Ca2+ crosstalk. Circ. Res. 111, 863–875 (2012).
Bassoy, E. Y. et al. ER-mitochondria contacts control surface glycan expression and sensitivity to killer lymphocytes in glioma stem-like cells. EMBO J. 36, 1493–1512 (2017).
Naon, D. et al. Does mitofusin 2 tether or separate endoplasmic reticulum and mitochondria? reply. Proc. Natl Acad. Sci. USA 114, E2268–E2269 (2017).
Cosson, P., Marchetti, A., Ravazzola, M. & Orci, L. Mitofusin-2 independent juxtaposition of endoplasmic reticulum and mitochondria: an ultrastructural study. PLoS One 7, e46293 (2012).
Filadi, R. et al. Mitofusin 2 ablation increases endoplasmic reticulum-mitochondria coupling. Proc. Natl Acad. Sci. USA 112, E2174–E2181 (2015).
Bravo, R. et al. Increased ER-mitochondrial coupling promotes mitochondrial respiration and bioenergetics during early phases of ER stress. J. Cell Sci. 124, 2143–2121s2152 (2011).
De Vos, K. J. et al. VAPB interacts with the mitochondrial protein PTPIP51 to regulate calcium homeostasis. Hum. Mol. Genet. 21, 1299–1311 (2012).
Gomez-Suaga, P. et al. The ER-mitochondria tethering complex VAPB-PTPIP51 regulates autophagy. Curr. Biol. 27, 371–385 (2017).
Kirmiz, M., Vierra, N. C., Palacio, S. & Trimmer, J. S. Identification of VAPA and VAPB as Kv2 channel-interacting proteins defining endoplasmic reticulum-plasma membrane junctions in mammalian brain neurons. J. Neurosci. 38, 7562–7584 (2018).
Hamasaki, M. et al. Autophagosomes form at ER-mitochondria contact sites. Nature 495, 389–393 (2013).
Cerqua, C. et al. Trichoplein/mitostatin regulates endoplasmic reticulum-mitochondria juxtaposition. EMBO Rep. 11, 854–860 (2010).
Dadsena, S. et al. Ceramides bind VDAC2 to trigger mitochondrial apoptosis. Nat. Commun. 10, 1832 (2019).
Soubannier, V. et al. A vesicular transport pathway shuttles cargo from mitochondria to lysosomes. Curr. Biol. 22, 135–141 (2012).
Magni, G., Emanuelli, M., Amici, A., Raffaelli, N. & Ruggieri, S. Purification of human nicotinamide-mononucleotide adenylyltransferase. Methods Enzymol. 280, 241–247 (1997).
Soubannier, V., Rippstein, P., Kaufman, B. A., Shoubridge, E. A. & McBride, H. M. Reconstitution of mitochondria derived vesicle formation demonstrates selective enrichment of oxidized cargo. PLoS One 7, e52830 (2012).
McLelland, G. L., Soubannier, V., Chen, C. X., McBride, H. M. & Fon, E. A. Parkin and PINK1 function in a vesicular trafficking pathway regulating mitochondrial quality control. EMBO J. 33, 282–295 (2014).
Waterham, H. R. et al. A lethal defect of mitochondrial and peroxisomal fission. N. Engl. J. Med. 356, 1736–1741 (2007).
Gerber, S. et al. Mutations in DNM1L, as in OPA1, result in dominant optic atrophy despite opposite effects on mitochondrial fusion and fission. Brain 140, 2586–2596 (2017).
Sheffer, R. et al. Postnatal microcephaly and pain insensitivity due to a de novo heterozygous DNM1L mutation causing impaired mitochondrial fission and function. Am. J. Med. Genet. A 170, 1603–1607 (2016).
Baxter, R. V. et al. Ganglioside-induced differentiation-associated protein-1 is mutant in Charcot-Marie-Tooth disease type 4A/8q21. Nat. Genet. 30, 21–22 (2002).
Boyer, O. et al. INF2 mutations in Charcot–Marie–Tooth disease with glomerulopathy. N. Engl. J. Med. 365, 2377–2388 (2011).
Brown, E. J. et al. Mutations in the formin gene INF2 cause focal segmental glomerulosclerosis. Nat. Genet. 42, 72 (2009).
Koch, J. et al. Disturbed mitochondrial and peroxisomal dynamics due to loss of MFF causes Leigh-like encephalopathy, optic atrophy and peripheral neuropathy. J. Med. Genet. 53, 270–278 (2016).
Shamseldin, H. E. et al. Genomic analysis of mitochondrial diseases in a consanguineous population reveals novel candidate disease genes. J. Med. Genet. 49, 234–241 (2012).
Schaaf, C. P. et al. Early-onset severe neuromuscular phenotype associated with compound heterozygosity for OPA1 mutations. Mol. Genet. Metab. 103, 383–387 (2011).
Spiegel, R. et al. Fatal infantile mitochondrial encephalomyopathy, hypertrophic cardiomyopathy and optic atrophy associated with a homozygous OPA1 mutation. J. Med. Genet. 53, 127–131 (2016).
Carelli, V. et al. Syndromic parkinsonism and dementia associated with OPA1 missense mutations. Ann. Neurol. 78, 21–38 (2015).
Harder, Z., Zunino, R. & McBride, H. Sumo1 conjugates mitochondrial substrates and participates in mitochondrial fission. Curr. Biol. 14, 340–345 (2004).
Yonashiro, R. et al. A novel mitochondrial ubiquitin ligase plays a critical role in mitochondrial dynamics. EMBO J. 25, 3618–3626 (2006).
Burchell, V. S. et al. The Parkinson’s disease–linked proteins Fbxo7 and parkin interact to mediate mitophagy. Nat. Neurosci. 16, 1257 (2013).
Park, Y.-Y. et al. Loss of MARCH5 mitochondrial E3 ubiquitin ligase induces cellular senescence through dynamin-related protein 1 and mitofusin 1. J. Cell Sci. 123, 619–626 (2010).
Pyakurel, A., Savoia, C., Hess, D. & Scorrano, L. Extracellular regulated kinase phosphorylates mitofusin 1 to control mitochondrial morphology and apoptosis. Mol. Cell 58, 244–254 (2015).
Zhou, W. et al. Mutation of the protein kinase a phosphorylation site influences the anti-proliferative activity of mitofusin 2. Atherosclerosis 211, 216–223 (2010).
Nakamura, N., Kimura, Y., Tokuda, M., Honda, S. & Hirose, S. MARCH-V is a novel mitofusin 2- and Drp1-binding protein able to change mitochondrial morphology. EMBO Rep. 7, 1019–1022 (2006).
Song, Z., Chen, H., Fiket, M., Alexander, C. & Chan, D. C. OPA1 processing controls mitochondrial fusion and is regulated by mRNA splicing, membrane potential, and Yme1L. J. Cell Biol. 178, 749–755 (2007).
Research in the L.S. laboratory is supported by AIRC IG19991, Italian Ministry of Education, University and Research PRIN 2017BF3PXZ, Fondation Leducq TNE15004, Muscular Dystrophy Association RG 603731 and Cariparo Foundation SIGMI. Research in the M.G. laboratory is supported by CARIPARO Starting Grant 2016 AIFbiol and Unipd STARS Consolidator FIRMESs.
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
- Krebs cycle
A series of chemical reactions catalysed by enzymes that, on oxidation of the acetyl portion of acetyl coenzyme A, generate reducing equivalents (NADH and FADH2).
- Membrane contact sites
Points at which two biological membranes run in parallel, at a constant distance, for several nanometres.
- Oxidative phosphorylation
(OXPHOS). A process involving coupling between mitochondrial respiration and ATP production.
- Mitochondrial respiratory chain
An ensemble of protein complexes whose coordinated activity catalyses the oxidation of reducing equivalents using oxygen as a terminal electron acceptor.
- F1Fo-ATP synthase
Protein complex located at the inner mitochondrial membrane which mediates Pi addition to ADP to generate ATP in a process driven by the passage of protons.
- Reducing equivalents
Electron donors in a redox reaction, which become oxidized. In mitochondria, NADH and FADH2.
- BCL-2 family
A group of evolutionarily conserved proteins that harbour a BCL-2 homology domain. Mostly known for their regulatory role in programmed cell death.
The literal meaning of this term is ‘nucleus-like’; it refers to a cluster of proteins and mitochondrial DNA, not surrounded by a membrane.
- Bin/amphiphysin/Rvs (BAR) domain
Domains that mediate protein dimerization, first discovered in three protein classes named Bin, amphiphysin and Rvs (BAR).
- Rhomboid protease
Intramembrane serine proteases that share a common catalytic domain composed of six membrane-spanning segments.
- Iron–sulphur clusters
Metal prosthetic groups synthesized in mitochondria and acting as cofactors to catalyse several redox enzymatic activities.
An iron-binding glycoprotein that controls the level of free iron in the blood.
The catabolic process through which fatty acids are degraded in mitochondria to produce acetyl coenzyme A.
A family of lipid molecules composed of a fatty acid conjugated to the 18-carbon amino alcohol sphingosine.
- Ischaemia–reperfusion injury
Tissue damage occurring especially in the heart on coronary artery occlusion and reperfusion on physicians’ intervention, for example, by percutaneous coronary intervention (formerly known as angioplasty with stent).
Omega-shaped organelles which precede the formation of autophagosomes.
- Retromer complex
A coat-like complex that mediates the recycling of the endosomal cargo into vesicles moving back to the Golgi apparatus.
- 5′-AMP-activated protein kinase
(AMPK). A key protein kinase that plays a critical role in cellular homeostasis. By phosphorylating a plethora of targets, it acts as a general switch to regulate cellular functions such as glucose absorption, fat oxidation and mitochondrial biogenesis.
- Chemiosmotic theory
The theory proposed by Mitchell explaining ATP synthesis by mitochondria. The chemical energy of NADH and FADH2 is converted by the electron-transport chain into an electrochemical proton gradient across the inner mitochondrial membrane (IMM), whose free energy is in turn used to catalyse the phosphorylation of ADP by the IMM-bound ATP synthase.
- Respiratory chain supercomplexes
Assembly of distinct respiratory chain complexes that guarantees efficient ATP production while reducing electron loss and production of reactive oxygen species.
- Epithelial–mesenchymal transition
The process through which epithelial cells lose their characteristics of adhesion and polarity and acquire those typical of mesenchymal stem cells, including differentiation abilities
- Nuclear factor of activated T cells
(NFAT). A family of transcription factors that can translocate into the nucleus in response to stimuli that enhance intracellular Ca2+ levels. Once in the nucleus, they induce transcription of target genes.
- Naive embryonic stem cells
Embryonic stem cells of the preimplantation blastocyst that have the potential to generate all the somatic cell lineages.
- Primed embryonic stem cells
Embryonic stem cells (from post-implantation epiblast) that can self-renew and differentiate.
- Senescence-associated secretory phenotype
The release of factors such as inflammatory cytokines, growth factors and proteases from senescent cells.
- Integrated stress response
A generally prosurvival stress response induced on exposure of the cell to either internal factors such as accumulation of unfolded protein leading to endoplasmic reticulum stress or external factors such as nutrient deprivation and viral infection.
Cyclic AMP-dependent transcription factor 4 plays a key role in cell response to stress stimuli, mostly by promoting cell survival.
A cytokine produced by tissues undergoing mitochondrial stress and affecting other cell types in an endocrine or a paracrine manner. Mitokines include, for example, FGF21 and GDF15.
This term derives from Greek leptos (‘thin’); it refers to a hormone that inhibits hunger.
- Pro-opiomelanocortin (POMC) neurons
Neurons located in the arcuate nucleus within the hypothalamus; they produce a polypeptide named pro-opiomelanocortin, whose proteolysis generates several peptide hormones (including α-melanocyte-stimulating hormone and adrenocorticotropic hormone).
- ER stress
A set of intracellular pathways activated by the cell to cope with abnormal accumulation of unfolded proteins in the lumen of the endoplasmic reticulum (ER).
- Brown adipocytes
Cells that compose the brown adipose tissue, enriched in mitochondria and characterized by high thermogenic potential.
- Alveolar type 2 epithelial cells
One of the two cell types that build the alveolar epithelium, responsible for surfactant synthesis and secretion.
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Giacomello, M., Pyakurel, A., Glytsou, C. et al. The cell biology of mitochondrial membrane dynamics. Nat Rev Mol Cell Biol 21, 204–224 (2020). https://doi.org/10.1038/s41580-020-0210-7
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