Skip to main content

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

  • Review Article
  • Published:

Energy matters: presynaptic metabolism and the maintenance of synaptic transmission

Abstract

Synaptic activity imposes large energy demands that are met by local adenosine triphosphate (ATP) synthesis through glycolysis and mitochondrial oxidative phosphorylation. ATP drives action potentials, supports synapse assembly and remodelling, and fuels synaptic vesicle filling and recycling, thus sustaining synaptic transmission. Given their polarized morphological features — including long axons and extensive branching in their terminal regions — neurons face exceptional challenges in maintaining presynaptic energy homeostasis, particularly during intensive synaptic activity. Recent studies have started to uncover the mechanisms and signalling pathways involved in activity-dependent and energy-sensitive regulation of presynaptic energetics, or ‘synaptoenergetics’. These conceptual advances have established the energetic regulation of synaptic efficacy and plasticity as an exciting research field that is relevant to a range of neurological disorders associated with bioenergetic failure and synaptic dysfunction.

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

Access options

Rent or buy this article

Prices vary by article type

from$1.95

to$39.95

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

Fig. 1: ATP generation and consumption at presynaptic terminals.
Fig. 2: Activity and Ca2+ signalling arrest motile mitochondria at presynaptic terminals.
Fig. 3: AMPK signalling regulates synaptoenergetics.
Fig. 4: Energetic regulation of synaptic transmission.

Similar content being viewed by others

References

  1. Kiser, B. Early child development: body of knowledge. Nature 523, 286–289 (2015).

    PubMed  Google Scholar 

  2. Chapman, E. R. A Ca2+ sensor for exocytosis. Trends Neurosci. 41, 327–330 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. Südhof, T. C. Towards an understanding of synapse formation. Neuron 100, 276–293 (2018).

    PubMed  PubMed Central  Google Scholar 

  4. Harris, J. J., Jolivet, R. & Attwell, D. Synaptic energy use and supply. Neuron 75, 762–777 (2012). This review provides an in-depth discussion of the consumption and supply of synaptic energy during development and synaptic plasticity and between wake and sleep states, and describes how defects in synaptic energy supply can lead to disease.

    CAS  PubMed  Google Scholar 

  5. Yellen, G. Fueling thought: management of glycolysis and oxidative phosphorylation in neuronal metabolism. J. Cell Biol. 217, 2235–2246 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  6. Attwell, D. & Laughlin, S. B. An energy budget for signaling in the grey matter of the brain. J. Cereb. Blood Flow. Metab. 21, 1133–1145 (2001).

    CAS  PubMed  Google Scholar 

  7. Magistretti, P. J. & Allaman, I. A cellular perspective on brain energy metabolism and functional imaging. Neuron 86, 883–901 (2015).

    CAS  PubMed  Google Scholar 

  8. Devine, M. J. & Kittler, J. T. Mitochondria at the neuronal presynapse in health and disease. Nat. Rev. Neurosci. 19, 63–80 (2018). This review discusses the importance of mitochondrial transport and positioning in rescaling presynaptic Ca2+ transients and homeostatic plasticity.

    CAS  PubMed  Google Scholar 

  9. Misgeld, T. & Schwarz, T. L. Mitostasis in neurons: maintaining mitochondria in an extended cellular architecture. Neuron 96, 651–666 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  10. Sheng, Z. H. Mitochondrial trafficking and anchoring in neurons: new insight and implications. J. Cell Biol. 204, 1087–1098 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  11. Sheng, Z. H. & Cai, Q. Mitochondrial transport in neurons: impact on synaptic homeostasis and neurodegeneration. Nat. Rev. Neurosci. 13, 77–93 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Mishra, P. & Chan, D. C. Metabolic regulation of mitochondrial dynamics. J. Cell Biol. 212, 379–387 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Ashrafi, G. & Ryan, T. A. Glucose metabolism in nerve terminals. Curr. Opin. Neurobiol. 45, 156–161 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Cunnane, S. C. et al. Brain energy rescue: an emerging therapeutic concept for neurodegenerative disorders of ageing. Nat. Rev. Drug Discov. 19, 609–633 (2020). This review discusses the status and prospect of therapeutic strategies for restoring glycolysis and oxidative phosphorylation in neurodegenerative disorders of ageing.

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Pathak, D., Berthet, A. & Nakamura, K. Energy failure: does it contribute to neurodegeneration? Ann. Neurol. 74, 506–516 (2013).

    PubMed  PubMed Central  Google Scholar 

  16. Mink, J. W., Blumenschine, R. J. & Adams, D. B. Ratio of central nervous system to body metabolism in vertebrates: its constancy and functional basis. Am. J. Physiol. 241, R203–R212 (1981).

    CAS  PubMed  Google Scholar 

  17. Harris, J. J. & Attwell, D. The energetics of CNS white matter. J. Neurosci. 32, 356–371 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  18. Rangaraju, V., Calloway, N. & Ryan, T. A. Activity-driven local ATP synthesis is required for synaptic function. Cell 156, 825–835 (2014). This study reports that synaptic activity drives local ATP synthesis through glycolysis and oxidative phosphorylation to meet presynaptic energy demands and that blocking glycolysis inhibits synaptic vesicle recycling.

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Schotten, S. et al. Additive effects on the energy barrier for synaptic vesicle fusion cause supralinear effects on the vesicle fusion rate. eLife 4, e05531 (2015).

    PubMed  PubMed Central  Google Scholar 

  20. Hyder, F., Fulbright, R. K., Shulman, R. G. & Rothman, D. L. Glutamatergic function in the resting awake human brain is supported by uniformly high oxidative energy. J. Cereb. Blood Flow. Metab. 33, 339–347 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. Fox, P. T., Raichle, M. E., Mintun, M. A. & Dence, C. Nonoxidative glucose consumption during focal physiologic neural activity. Science 241, 462–464 (1988).

    CAS  PubMed  Google Scholar 

  22. Dhar-Chowdhury, P., Malester, B., Rajacic, P. & Coetzee, W. A. The regulation of ion channels and transporters by glycolytically derived ATP. Cell Mol. Life Sci. 64, 3069–3083 (2007).

    CAS  PubMed  Google Scholar 

  23. Hinckelmann, M. V. et al. Self-propelling vesicles define glycolysis as the minimal energy machinery for neuronal transport. Nat. Commun. 7, 13233 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Ikemoto, A., Bole, D. G. & Ueda, T. Glycolysis and glutamate accumulation into synaptic vesicles. Role of glyceraldehyde phosphate dehydrogenase and 3-phosphoglycerate kinase. J. Biol. Chem. 278, 5929–5940 (2003).

    CAS  PubMed  Google Scholar 

  25. Zala, D. et al. Vesicular glycolysis provides on-board energy for fast axonal transport. Cell 152, 479–491 (2013).

    CAS  PubMed  Google Scholar 

  26. Sobieski, C., Fitzpatrick, M. J. & Mennerick, S. J. Differential presynaptic ATP supply for basal and high-demand transmission. J. Neurosci. 37, 1888–1899 (2017). This study shows that basal evoked synaptic transmission can be sustained by activity-boosted glycolysis, but that intensive synaptic activity requires oxidative phosphorylation.

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Rose, J., Brian, C., Pappa, A., Panayiotidis, M. I. & Franco, R. Mitochondrial metabolism in astrocytes regulates brain bioenergetics, neurotransmission and redox balance. Front. Neurosci. 14, 536682 (2020).

    PubMed  PubMed Central  Google Scholar 

  28. Hyder, F. et al. Neuronal-glial glucose oxidation and glutamatergic-GABAergic function. J. Cereb. Blood Flow Metab. 26, 865–877 (2006).

    CAS  PubMed  Google Scholar 

  29. Zhang, Y. et al. An RNA-sequencing transcriptome and splicing database of glia, neurons, and vascular cells of the cerebral cortex. J. Neurosci. 34, 11929–11947 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Pellerin, L. & Magistretti, P. J. Glutamate uptake into astrocytes stimulates aerobic glycolysis: a mechanism coupling neuronal activity to glucose utilization. Proc. Natl Acad. Sci. USA 91, 10625–10629 (1994). This pioneering study led to the proposal of the astrocyte–neuron lactate shuttle hypothesis.

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Allen, N. J., Káradóttir, R. & Attwell, D. A preferential role for glycolysis in preventing the anoxic depolarization of rat hippocampal area CA1 pyramidal cells. J. Neurosci. 25, 848–859 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Bak, L. K. et al. Neuronal glucose but not lactate utilization is positively correlated with NMDA-induced neurotransmission and fluctuations in cytosolic Ca2+ levels. J. Neurochem. 109, 87–93 (2009).

    CAS  PubMed  Google Scholar 

  33. Barros, L. F. & Weber, B. CrossTalk proposal: an important astrocyte-to-neuron lactate shuttle couples neuronal activity to glucose utilisation in the brain. J. Physiol. 596, 347–350 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Chih, C. P., Lipton, P. & Roberts, E. L. Do active cerebral neurons really use lactate rather than glucose? Trends Neurosci. 24, 573–578 (2001).

    CAS  PubMed  Google Scholar 

  35. Dienel, G. A. Lack of appropriate stoichiometry: strong evidence against an energetically important astrocyte-neuron lactate shuttle in brain. J. Neurosci. Res. 95, 2103–2125 (2017).

    CAS  PubMed  Google Scholar 

  36. Gjedde, A. & Marrett, S. Glycolysis in neurons, not astrocytes, delays oxidative metabolism of human visual cortex during sustained checkerboard stimulation in vivo. J. Cereb. Blood Flow Metab. 21, 1384–1392 (2001).

    CAS  PubMed  Google Scholar 

  37. Hall, C. N., Klein-Flügge, M. C., Howarth, C. & Attwell, D. Oxidative phosphorylation, not glycolysis, powers presynaptic and postsynaptic mechanisms underlying brain information processing. J. Neurosci. 32, 8940–8951 (2012). This study characterizes the brain energy budget and shows that synaptic function is powered mainly by oxidative phosphorylation.

    CAS  PubMed  PubMed Central  Google Scholar 

  38. Hertz, L. The astrocyte-neuron lactate shuttle: a challenge of a challenge. J. Cereb. Blood Flow Metab. 24, 1241–1248 (2004).

    PubMed  Google Scholar 

  39. Murphy-Royal, C. et al. Stress gates an astrocytic energy reservoir to impair synaptic plasticity. Nat. Commun. 11, 2014 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Bouzier-Sore, A. K. et al. Competition between glucose and lactate as oxidative energy substrates in both neurons and astrocytes: a comparative NMR study. Eur. J. Neurosci. 24, 1687–1694 (2006).

    PubMed  Google Scholar 

  41. Lange, S. C. et al. Dynamic changes in cytosolic ATP levels in cultured glutamatergic neurons during NMDA-induced synaptic activity supported by glucose or lactate. Neurochem. Res. 40, 2517–2526 (2015).

    CAS  PubMed  Google Scholar 

  42. Lundgaard, I. et al. Direct neuronal glucose uptake heralds activity-dependent increases in cerebral metabolism. Nat. Commun. 6, 6807 (2015).

    CAS  PubMed  Google Scholar 

  43. Lucas, S. J. et al. Glucose and lactate as metabolic constraints on presynaptic transmission at an excitatory synapse. J. Physiol. 596, 1699–1721 (2018). This study shows that presynaptic ATP depletion at the calyx of Held accelerates synaptic depression during HFS owing to the failed replenishment of the readily releasable vesicle pool.

    CAS  PubMed  PubMed Central  Google Scholar 

  44. Hollnagel, J. O. et al. Lactate attenuates synaptic transmission and affects brain rhythms featuring high energy expenditure. iScience 23, 101316 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Díaz-García, C. M. et al. Neuronal stimulation triggers neuronal glycolysis and not lactate uptake. Cell Metab. 26, 361–374.e4 (2017). This study shows that neuronal activity transiently increases the cytosolic NADH/NAD+ ratio by enhancing glucose consumption in neurons, but not in astrocytes.

    PubMed  PubMed Central  Google Scholar 

  46. Dubinsky, W. P., Mayorga-Wark, O. & Schultz, S. G. Colocalization of glycolytic enzyme activity and KATP channels in basolateral membrane of Necturus enterocytes. Am. J. Physiol. 275, C1653–C1659 (1998).

    CAS  PubMed  Google Scholar 

  47. Mercer, R. W. & Dunham, P. B. Membrane-bound ATP fuels the Na/K pump. Studies on membrane-bound glycolytic enzymes on inside-out vesicles from human red cell membranes. J. Gen. Physiol. 78, 547–568 (1981).

    CAS  PubMed  Google Scholar 

  48. Lu, M., Holliday, L. S., Zhang, L., Dunn, W. A. Jr & Gluck, S. L. Interaction between aldolase and vacuolar H+-ATPase: evidence for direct coupling of glycolysis to the ATP-hydrolyzing proton pump. J. Biol. Chem. 276, 30407–30413 (2001).

    CAS  PubMed  Google Scholar 

  49. Paul, R. J., Hardin, C. D., Raeymaekers, L., Wuytack, F. & Casteels, R. Preferential support of Ca2+ uptake in smooth muscle plasma membrane vesicles by an endogenous glycolytic cascade. FASEB J. 3, 2298–2301 (1989).

    CAS  PubMed  Google Scholar 

  50. Lujan, B., Kushmerick, C., Banerjee, T. D., Dagda, R. K. & Renden, R. Glycolysis selectively shapes the presynaptic action potential waveform. J. Neurophysiol. 116, 2523–2540 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  51. Wang, P. et al. A Drosophila temperature-sensitive seizure mutant in phosphoglycerate kinase disrupts ATP generation and alters synaptic function. J. Neurosci. 24, 4518–4529 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Knull, H. R. Compartmentation of glycolytic enzymes in nerve endings as determined by glutaraldehyde fixation. J. Biol. Chem. 255, 6439–6444 (1980).

    CAS  PubMed  Google Scholar 

  53. Jang, S. et al. Glycolytic enzymes localize to synapses under energy stress to support synaptic function. Neuron 90, 278–291 (2016). This study shows that the glycolytic metabolon is redistributed to synapses under energy stress to sustain synaptic function and behaviour in worm neurons.

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Gerhart, D. Z., Broderius, M. A., Borson, N. D. & Drewes, L. R. Neurons and microvessels express the brain glucose transporter protein GLUT3. Proc. Natl Acad. Sci. USA 89, 733–737 (1992).

    CAS  PubMed  PubMed Central  Google Scholar 

  55. Ferreira, J. M., Burnett, A. L. & Rameau, G. A. Activity-dependent regulation of surface glucose transporter-3. J. Neurosci. 31, 1991–1999 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  56. Kobayashi, M., Nikami, H., Morimatsu, M. & Saito, M. Expression and localization of insulin-regulatable glucose transporter (GLUT4) in rat brain. Neurosci. Lett. 213, 103–106 (1996).

    CAS  PubMed  Google Scholar 

  57. Pearson-Leary, J. & McNay, E. C. Novel roles for the insulin-regulated glucose transporter-4 in hippocampally dependent memory. J. Neurosci. 36, 11851–11864 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  58. Ashrafi, G., Wu, Z., Farrell, R. J. & Ryan, T. A. GLUT4 mobilization supports energetic demands of active synapses. Neuron 93, 606–615.e3 (2017). This study reveals AMPK-dependent translocation of GLUT4 into the presynaptic membrane to meet activity-driven increases in energy demand.

    CAS  PubMed  PubMed Central  Google Scholar 

  59. Douen, A. G. et al. Exercise induces recruitment of the “insulin-responsive glucose transporter”. Evidence for distinct intracellular insulin- and exercise-recruitable transporter pools in skeletal muscle. J. Biol. Chem. 265, 13427–13430 (1990).

    CAS  PubMed  Google Scholar 

  60. Koepsell, H. Glucose transporters in brain in health and disease. Pflug. Arch. 472, 1299–1343 (2020).

    CAS  Google Scholar 

  61. Pekkurnaz, G., Trinidad, J. C., Wang, X., Kong, D. & Schwarz, T. L. Glucose regulates mitochondrial motility via Milton modification by O-GlcNAc transferase. Cell 158, 54–68 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. Pfeiffer, T., Schuster, S. & Bonhoeffer, S. Cooperation and competition in the evolution of ATP-producing pathways. Science 292, 504–507 (2001).

    CAS  PubMed  Google Scholar 

  63. Pathak, D. et al. The role of mitochondrially derived ATP in synaptic vesicle recycling. J. Biol. Chem. 290, 22325–22336 (2015). This study shows that mitochondrion-derived ATP powers synaptic vesicle endocytosis and recycling at presynaptic boutons.

    CAS  PubMed  PubMed Central  Google Scholar 

  64. Davis, A. F. & Clayton, D. A. In situ localization of mitochondrial DNA replication in intact mammalian cells. J. Cell Biol. 135, 883–893 (1996).

    CAS  PubMed  Google Scholar 

  65. Amiri, M. & Hollenbeck, P. J. Mitochondrial biogenesis in the axons of vertebrate peripheral neurons. Dev. Neurobiol. 68, 1348–1361 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. Van Laar, V. S. et al. Evidence for compartmentalized axonal mitochondrial biogenesis: mitochondrial DNA replication increases in distal axons as an early response to Parkinson’s disease-relevant stress. J. Neurosci. 38, 7505–7515 (2018).

    PubMed  PubMed Central  Google Scholar 

  67. Kuzniewska, B. et al. Mitochondrial protein biogenesis in the synapse is supported by local translation. EMBO Rep. 21, e48882 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. LeBleu, V. S. et al. PGC-1alpha mediates mitochondrial biogenesis and oxidative phosphorylation in cancer cells to promote metastasis. Nat. Cell Biol. 16, 992–1003 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Lin, T. H. et al. TSG101 negatively regulates mitochondrial biogenesis in axons. Proc. Natl Acad. Sci. USA 118, e2018770118 (2021).

    CAS  PubMed  PubMed Central  Google Scholar 

  70. Chen, H. & Chan, D. C. Mitochondrial dynamics — fusion, fission, movement, and mitophagy — in neurodegenerative diseases. Hum. Mol. Genet. 18, R169–R176 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  71. 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).

    CAS  PubMed  Google Scholar 

  72. Berthet, A. et al. Loss of mitochondrial fission depletes axonal mitochondria in midbrain dopamine neurons. J. Neurosci. 34, 14304–14317 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. Shields, L. Y. et al. Dynamin-related protein 1 is required for normal mitochondrial bioenergetic and synaptic function in CA1 hippocampal neurons. Cell Death Dis. 6, e1725 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  74. Oettinghaus, B. et al. Synaptic dysfunction, memory deficits and hippocampal atrophy due to ablation of mitochondrial fission in adult forebrain neurons. Cell Death Differ. 23, 18–28 (2016).

    CAS  PubMed  Google Scholar 

  75. Verstreken, P. et al. Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions. Neuron 47, 365–378 (2005). This study shows that, in the Drosophila Drp1 mutant, failed mitochondrial localization at neuromuscular junctions results in faster synaptic vesicle depletion under HFS, due to defective synaptic vesicle mobilization from the reverse pool.

    CAS  PubMed  Google Scholar 

  76. Lewis, T. L., Kwon, S. K., Lee, A., Shaw, R. & Polleux, F. MFF-dependent mitochondrial fission regulates presynaptic release and axon branching by limiting axonal mitochondria size. Nat. Commun. 9, 5008 (2018).

    PubMed  PubMed Central  Google Scholar 

  77. Singh, M., Denny, H., Smith, C., Granados, J. & Renden, R. Presynaptic loss of dynamin-related protein 1 impairs synaptic vesicle release and recycling at the mouse calyx of Held. J. Physiol. 596, 6263–6287 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. Song, Z., Ghochani, M., McCaffery, J. M., Frey, T. G. & Chan, D. C. Mitofusins and OPA1 mediate sequential steps in mitochondrial membrane fusion. Mol. Biol. Cell 20, 3525–3532 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. Iannielli, A. et al. Reconstitution of the human nigro-striatal pathway on-a-chip reveals OPA1-dependent mitochondrial defects and loss of dopaminergic synapses. Cell Rep. 29, 4646–4656 e4644 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Baloh, R. H., Schmidt, R. E., Pestronk, A. & Milbrandt, J. Altered axonal mitochondrial transport in the pathogenesis of Charcot-Marie-Tooth disease from mitofusin 2 mutations. J. Neurosci. 27, 422–430 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  81. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  82. Narendra, D. P. et al. PINK1 is selectively stabilized on impaired mitochondria to activate Parkin. PLoS Biol. 8, e1000298 (2010).

    PubMed  PubMed Central  Google Scholar 

  83. Devireddy, S., Liu, A., Lampe, T. & Hollenbeck, P. J. The organization of mitochondrial quality control and life cycle in the nervous system in vivo in the absence of PINK1. J. Neurosci. 35, 9391–9401 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. Lin, M. Y. et al. Releasing syntaphilin removes stressed mitochondria from axons independent of mitophagy under pathophysiological conditions. Neuron 94, 595–610.e6 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Sung, H., Tandarich, L. C., Nguyen, K. & Hollenbeck, P. J. Compartmentalized regulation of Parkin-mediated mitochondrial quality control in the Drosophila nervous system in vivo. J. Neurosci. 36, 7375–7391 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  86. Ashrafi, G., Schlehe, J. S., LaVoie, M. J. & Schwarz, T. L. Mitophagy of damaged mitochondria occurs locally in distal neuronal axons and requires PINK1 and Parkin. J. Cell Biol. 206, 655–670 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  87. Cai, Q., Zakaria, H. M., Simone, A. & Sheng, Z. H. Spatial parkin translocation and degradation of damaged mitochondria via mitophagy in live cortical neurons. Curr. Biol. 22, 545–552 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. Puri, R., Cheng, X. T., Lin, M. Y., Huang, N. & Sheng, Z. H. Mul1 restrains Parkin-mediated mitophagy in mature neurons by maintaining ER-mitochondrial contacts. Nat. Commun. 10, 3645 (2019).

    PubMed  PubMed Central  Google Scholar 

  89. Van Laar, V. S. et al. Bioenergetics of neurons inhibit the translocation response of Parkin following rapid mitochondrial depolarization. Hum. Mol. Genet. 20, 927–940 (2011).

    PubMed  Google Scholar 

  90. Sun, T., Qiao, H., Pan, P. Y., Chen, Y. & Sheng, Z. H. Motile axonal mitochondria contribute to the variability of presynaptic strength. Cell Rep. 4, 413–419 (2013). This study shows that mobile mitochondria passing through presynaptic boutons cause wide fluctuations in local ATP levels and thus contribute to pulse-to-pulse presynaptic variability.

    CAS  PubMed  PubMed Central  Google Scholar 

  91. Kang, J. S. et al. Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation. Cell 132, 137–148 (2008). This study reveals SNPH to be a ‘static anchor’ that selectively holds axonal mitochondria stationary via its docking interaction with microtubules.

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Li, S., Xiong, G. J., Huang, N. & Sheng, Z. H. The cross-talk of energy sensing and mitochondrial anchoring sustains synaptic efficacy by maintaining presynaptic metabolism. Nat. Metab. 2, 1077–1095 (2020). This study reveals crosstalk between AMPK–PAK energy sensing and SNPH–MYO6-mediated mitochondrial anchoring that sustains prolonged synaptic efficacy.

    CAS  PubMed  PubMed Central  Google Scholar 

  93. Shulman, Y. et al. ATP binding to synaspsin IIa regulates usage and clustering of vesicles in terminals of hippocampal neurons. J. Neurosci. 35, 985–998 (2015).

    PubMed  PubMed Central  Google Scholar 

  94. Smith, H. L. et al. Mitochondrial support of persistent presynaptic vesicle mobilization with age-dependent synaptic growth after LTP. eLife 5, e15275 (2016). This three-dimensional electron microscopy study shows that only about 33% of presynaptic active zones in hippocampi retain mitochondria and that sustained synaptic activity is restricted to mitochondrion-containing active zones.

    PubMed  PubMed Central  Google Scholar 

  95. Graffe, M., Zenisek, D. & Taraska, J. W. A marginal band of microtubules transports and organizes mitochondria in retinal bipolar synaptic terminals. J. Gen. Physiol. 146, 109–117 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. Perkins, G. A. et al. The micro-architecture of mitochondria at active zones: electron tomography reveals novel anchoring scaffolds and cristae structured for high-rate metabolism. J. Neurosci. 30, 1015–1026 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. Kamer, K. J. & Mootha, V. K. The molecular era of the mitochondrial calcium uniporter. Nat. Rev. Mol. Cell Biol. 16, 545–553 (2015).

    CAS  PubMed  Google Scholar 

  98. Duchen, M. R. Ca2+-dependent changes in the mitochondrial energetics in single dissociated mouse sensory neurons. Biochem. J. 283, 41–50 (1992).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. Jouaville, L. S., Pinton, P., Bastianutto, C., Rutter, G. A. & Rizzuto, R. Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic priming. Proc. Natl Acad. Sci. USA 96, 13807–13812 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  100. Llorente-Folch, I. et al. The regulation of neuronal mitochondrial metabolism by calcium. J. Physiol. 593, 3447–3462 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. McCormack, J. G., Halestrap, A. P. & Denton, R. M. Role of calcium ions in regulation of mammalian intramitochondrial metabolism. Physiol. Rev. 70, 391–425 (1990).

    CAS  PubMed  Google Scholar 

  102. Balaban, R. S. Cardiac energy metabolism homeostasis: role of cytosolic calcium. J. Mol. Cell Cardiol. 34, 1259–1271 (2002).

    CAS  PubMed  Google Scholar 

  103. Ashrafi, G., de Juan-Sanz, J., Farrell, R. J. & Ryan, T. A. Molecular tuning of the axonal mitochondrial Ca2+ uniporter ensures metabolic flexibility of neurotransmission. Neuron 105, 678–687.e5 (2020).

    CAS  PubMed  Google Scholar 

  104. Chouhan, A. K. et al. Cytosolic calcium coordinates mitochondrial energy metabolism with presynaptic activity. J. Neurosci. 32, 1233–1243 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Alavian, K. N. et al. Bcl-xL regulates metabolic efficiency of neurons through interaction with the mitochondrial F1FO ATP synthase. Nat. Cell Biol. 13, 1224–1233 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  106. Li, H. et al. Bcl-xL induces Drp1-dependent synapse formation in cultured hippocampal neurons. Proc. Natl Acad. Sci. USA 105, 2169–2174 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  107. Berman, S. B. et al. Bcl-xL increases mitochondrial fission, fusion, and biomass in neurons. J. Cell Biol. 184, 707–719 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Perkins, G. A., Ellisman, M. H. & Fox, D. A. Three-dimensional analysis of mouse rod and cone mitochondrial cristae architecture: bioenergetic and functional implications. Mol. Vis. 9, 60–73 (2003).

    CAS  PubMed  Google Scholar 

  109. Cogliati, S. et al. Mitochondrial cristae shape determines respiratory chain supercomplexes assembly and respiratory efficiency. Cell 155, 160–171 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  110. Gomes, L. C., Di Benedetto, G. & Scorrano, L. During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Nielsen, J. et al. Plasticity in mitochondrial cristae density allows metabolic capacity modulation in human skeletal muscle. J. Physiol. 595, 2839–2847 (2017).

    CAS  PubMed  Google Scholar 

  112. Cserép, C., Pósfai, B., Schwarcz, A. D. & Dénes, A. Mitochondrial ultrastructure is coupled to synaptic performance at axonal release sites. eNeuro 5, https://doi.org/10.1523/ENEURO.0390-17.2018 (2018).

  113. Stauch, K. L. et al. Quantitative proteomics of presynaptic mitochondria reveal an overexpression and biological relevance of neuronal MitoNEET in postnatal brain development. Dev. Neurobiol. 79, 370–386 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  114. Völgyi, K. et al. Synaptic mitochondria: a brain mitochondria cluster with a specific proteome. J. Proteom. 120, 142–157 (2015).

    Google Scholar 

  115. Chang, D. T. & Reynolds, I. J. Mitochondrial trafficking and morphology in healthy and injured neurons. Prog. Neurobiol. 80, 241–268 (2006).

    CAS  PubMed  Google Scholar 

  116. Morris, R. L. & Hollenbeck, P. J. The regulation of bidirectional mitochondrial transport is coordinated with axonal outgrowth. J. Cell Sci. 104, 917–927 (1993).

    PubMed  Google Scholar 

  117. Morsci, N. S., Hall, D. H., Driscoll, M. & Sheng, Z. H. Age-related phasic patterns of mitochondrial maintenance in adult Caenorhabditis elegans neurons. J. Neurosci. 36, 1373–1385 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Ruthel, G. & Hollenbeck, P. J. Response of mitochondrial traffic to axon determination and differential branch growth. J. Neurosci. 23, 8618–8624 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. Vagnoni, A. & Bullock, S. L. A cAMP/PKA/kinesin-1 axis promotes the axonal transport of mitochondria in aging Drosophila neurons. Curr. Biol. 28, 1265–1272.e4 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  120. Zhou, B. et al. Facilitation of axon regeneration by enhancing mitochondrial transport and rescuing energy deficits. J. Cell Biol. 214, 103–119 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Lees, R. M., Johnson, J. D. & Ashby, M. C. Presynaptic boutons that contain mitochondria are more stable. Front. Synaptic Neurosci. 11, 37 (2020).

    PubMed  PubMed Central  Google Scholar 

  122. Misgeld, T., Kerschensteiner, M., Bareyre, F. M., Burgess, R. W. & Lichtman, J. W. Imaging axonal transport of mitochondria in vivo. Nat. Methods 4, 559–561 (2007). This study provides the first in vivo evidence of low mitochondrial motility in the adult mouse nervous system.

    CAS  PubMed  Google Scholar 

  123. Lewis, T. L., Turi, G. F., Kwon, S. K., Losonczy, A. & Polleux, F. Progressive decrease of mitochondrial motility during maturation of cortical axons in vitro and in vivo. Curr. Biol. 26, 2602–2608 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. Smit-Rigter, L. et al. Mitochondrial dynamics in visual cortex are limited in vivo and not affected by axonal structural plasticity. Curr. Biol. 26, 2609–2616 (2016).

    CAS  PubMed  Google Scholar 

  125. Saxton, W. M. & Hollenbeck, P. J. The axonal transport of mitochondria. J. Cell Sci. 125, 2095–2104 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  126. Sheng, Z. H. The interplay of axonal energy homeostasis and mitochondrial trafficking and anchoring. Trends Cell Biol. 27, 403–416 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. Chen, Y. & Sheng, Z. H. Kinesin-1-syntaphilin coupling mediates activity-dependent regulation of axonal mitochondrial transport. J. Cell Biol. 202, 351–364 (2013). This study shows that SNPH stably anchors axonal mitochondria by coordinating with MIRO1–Ca2+ signalling.

    CAS  PubMed  PubMed Central  Google Scholar 

  128. Rintoul, G. L., Filiano, A. J., Brocard, J. B., Kress, G. J. & Reynolds, I. J. Glutamate decreases mitochondrial size and movement in primary forebrain neurons. J. Neurosci. 23, 7881–7888 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  129. Saotome, M. et al. Bidirectional Ca2+-dependent control of mitochondrial dynamics by the Miro GTPase. Proc. Natl Acad. Sci. USA 105, 20728–20733 (2008). This study shows that MIRO1 arrests mitochondrial transport by sensing Ca2+.

    CAS  PubMed  PubMed Central  Google Scholar 

  130. MacAskill, A. F. et al. Miro1 is a calcium sensor for glutamate receptor-dependent localization of mitochondria at synapses. Neuron 61, 541–555 (2009). This is one of two studies, along with Wang and Schwarz (2009), showing that MIRO1–Ca2+ signalling arrests motile mitochondria at activated synapses.

    CAS  PubMed  PubMed Central  Google Scholar 

  131. Wang, X. & Schwarz, T. L. The mechanism of Ca2+-dependent regulation of kinesin-mediated mitochondrial motility. Cell 136, 163–174 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  132. Vaccaro, V., Devine, M. J., Higgs, N. F. & Kittler, J. T. Miro1-dependent mitochondrial positioning drives the rescaling of presynaptic Ca2+ signals during homeostatic plasticity. EMBO Rep. 18, 231–240 (2017). This study reports that MIRO1–Ca2+ sensing recruits presynaptic mitochondria to rescale the strength of synaptic transmission during homeostatic plasticity.

    CAS  PubMed  Google Scholar 

  133. Gutnick, A., Banghart, M. R., West, E. R. & Schwarz, T. L. The light-sensitive dimerizer zapalog reveals distinct modes of immobilization for axonal mitochondria. Nat. Cell Biol. 21, 768–777 (2019). This optogenetic study shows that inhibiting actin polymerization reduces anchored mitochondria at VGLUT1-positive presynapses.

    CAS  PubMed  PubMed Central  Google Scholar 

  134. Nguyen, T. T. et al. Loss of Miro1-directed mitochondrial movement results in a novel murine model for neuron disease. Proc. Natl Acad. Sci. USA 111, E3631–E3640 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  135. López-Doménech, G. et al. Loss of dendritic complexity precedes neurodegeneration in a mouse model with disrupted mitochondrial distribution in mature dendrites. Cell Rep. 17, 317–327 (2016).

    PubMed  PubMed Central  Google Scholar 

  136. López-Doménech, G. et al. Miro proteins coordinate microtubule- and actin-dependent mitochondrial transport and distribution. EMBO J. 37, 321–336 (2018). This study reports that TRAK1/2 can drive mitochondrial transport in Miro1Miro2-double-knockout cells and that MIRO–MYO19 fine-tunes mitochondrial trafficking and positioning.

    PubMed  PubMed Central  Google Scholar 

  137. Chen, S., Owens, G. C. & Edelman, D. B. Dopamine inhibits mitochondrial motility in hippocampal neurons. PLoS ONE 3, e2804 (2008).

    PubMed  PubMed Central  Google Scholar 

  138. Morris, R. L. & Hollenbeck, P. J. Axonal transport of mitochondria along microtubules and F-actin in living vertebrate neurons. J. Cell Biol. 131, 1315–1326 (1995). This study reveals that F-actin and microtubules mediate axonal mitochondrial transport with different velocities and properties.

    CAS  PubMed  Google Scholar 

  139. Rintoul, G. L., Bennett, V. J., Papaconstandinou, N. A. & Reynolds, I. J. Nitric oxide inhibits mitochondrial movement in forebrain neurons associated with disruption of mitochondrial membrane potential. J. Neurochem. 97, 800–806 (2006).

    CAS  PubMed  Google Scholar 

  140. Zanelli, S. A., Trimmer, P. A. & Solenski, N. J. Nitric oxide impairs mitochondrial movement in cortical neurons during hypoxia. J. Neurochem. 97, 724–736 (2006).

    CAS  PubMed  Google Scholar 

  141. Chada, S. R. & Hollenbeck, P. J. Nerve growth factor signaling regulates motility and docking of axonal mitochondria. Curr. Biol. 14, 1272–1276 (2004).

    CAS  PubMed  Google Scholar 

  142. Pathak, D., Sepp, K. J. & Hollenbeck, P. J. Evidence that myosin activity opposes microtubule-based axonal transport of mitochondria. J. Neurosci. 30, 8984–8992 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  143. Quintero, O. A. et al. Human Myo19 is a novel myosin that associates with mitochondria. Curr. Biol. 19, 2008–2013 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  144. Cingolani, L. A. & Goda, Y. Actin in action: the interplay between the actin cytoskeleton and synaptic efficacy. Nat. Rev. Neurosci. 9, 344–356 (2008).

    CAS  PubMed  Google Scholar 

  145. Coles, C. H. & Bradke, F. Coordinating neuronal actin-microtubule dynamics. Curr. Biol. 25, R677–R691 (2015).

    CAS  PubMed  Google Scholar 

  146. Xu, K., Zhong, G. & Zhuang, X. Actin, spectrin, and associated proteins form a periodic cytoskeletal structure in axons. Science 339, 452–456 (2013).

    CAS  PubMed  Google Scholar 

  147. Doménech, E. et al. AMPK and PFKFB3 mediate glycolysis and survival in response to mitophagy during mitotic arrest. Nat. Cell Biol. 17, 1304–1316 (2015).

    PubMed  Google Scholar 

  148. Herzig, S. & Shaw, R. J. AMPK: guardian of metabolism and mitochondrial homeostasis. Nat. Rev. Mol. Cell Biol. 19, 121–135 (2018).

    CAS  PubMed  Google Scholar 

  149. Kishton, R. J. et al. AMPK is essential to balance glycolysis and mitochondrial metabolism to control T-ALL cell stress and survival. Cell Metab. 23, 649–662 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  150. Marinangeli, C. et al. AMP-activated protein kinase is essential for the maintenance of energy levels during synaptic activation. iScience 9, 1–13 (2018). This study shows that AMPK maintains neuronal energy levels upon synaptic activation by adapting the rate of glycolysis and mitochondrial respiration.

    CAS  PubMed  PubMed Central  Google Scholar 

  151. Yu, D. F. et al. HFS-triggered AMPK activation phosphorylates GSK3β and induces E-LTP in rat hippocampus in vivo. CNS Neurosci. Ther. 22, 525–531 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Tao, K., Matsuki, N. & Koyama, R. AMP-activated protein kinase mediates activity-dependent axon branching by recruiting mitochondria to axon. Dev. Neurobiol. 74, 557–573 (2014). This study shows that AMPK activation enhances mitochondrial transport and facilitates mitochondrial distribution at presynaptic terminals.

    CAS  PubMed  Google Scholar 

  153. Yang, W., Zhou, X., Zimmermann, H. R. & Ma, T. Brain-specific suppression of AMPKα2 isoform impairs cognition and hippocampal LTP by PERK-mediated eIF2α phosphorylation. Mol. Psychiatry 26, 1880–1897 (2021).

    CAS  PubMed  Google Scholar 

  154. Watters, O., Connolly, N. M. C., König, H. G., Düssmann, H. & Prehn, J. H. M. AMPK preferentially depresses retrograde transport of axonal mitochondria during localized nutrient deprivation. J. Neurosci. 40, 4798–4812 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Courchet, J. et al. Terminal axon branching is regulated by the LKB1-NUAK1 kinase pathway via presynaptic mitochondrial capture. Cell 153, 1510–1525 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  156. Kong, D. et al. A postsynaptic AMPK—>p21-activated kinase pathway drives fasting-induced synaptic plasticity in AgRP neurons. Neuron 91, 25–33 (2016). This study shows that fasting activates AMPK in hypothalamic AGRP neurons, and that this activates PAK signalling to maintain AGRP neuron state-dependent plasticity.

    CAS  PubMed  PubMed Central  Google Scholar 

  157. Bokoch, G. M. Biology of the p21-activated kinases. Annu. Rev. Biochem. 72, 743–781 (2003).

    CAS  PubMed  Google Scholar 

  158. Meng, J., Meng, Y., Hanna, A., Janus, C. & Jia, Z. Abnormal long-lasting synaptic plasticity and cognition in mice lacking the mental retardation gene Pak3. J. Neurosci. 25, 6641–6650 (2005). This study finds that Pak3-knockout mice exhibit significant abnormalities in synaptic plasticity.

    CAS  PubMed  PubMed Central  Google Scholar 

  159. Murthy, V. N., Sejnowski, T. J. & Stevens, C. F. Heterogeneous release properties of visualized individual hippocampal synapses. Neuron 18, 599–612 (1997).

    CAS  PubMed  Google Scholar 

  160. Atwood, H. L. & Karunanithi, S. Diversification of synaptic strength: presynaptic elements. Nat. Rev. Neurosci. 3, 497–516 (2002).

    CAS  PubMed  Google Scholar 

  161. Marder, E. & Goaillard, J. M. Variability, compensation and homeostasis in neuron and network function. Nat. Rev. Neurosci. 7, 563–574 (2006).

    CAS  PubMed  Google Scholar 

  162. Branco, T. & Staras, K. The probability of neurotransmitter release: variability and feedback control at single synapses. Nat. Rev. Neurosci. 10, 373–383 (2009).

    CAS  PubMed  Google Scholar 

  163. Ribrault, C., Sekimoto, K. & Triller, A. From the stochasticity of molecular processes to the variability of synaptic transmission. Nat. Rev. Neurosci. 12, 375–387 (2011).

    CAS  PubMed  Google Scholar 

  164. Billups, B. & Forsythe, I. D. Presynaptic mitochondrial calcium sequestration influences transmission at mammalian central synapses. J. Neurosci. 22, 5840–5847 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  165. Friel, D. D. & Tsien, R. W. An FCCP-sensitive Ca2+ store in bullfrog sympathetic neurons and its participation in stimulus-evoked changes in [Ca2+]i. J. Neurosci. 14, 4007–4024 (1994).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. Tang, Y. & Zucker, R. S. Mitochondrial involvement in post-tetanic potentiation of synaptic transmission. Neuron 18, 483–491 (1997).

    CAS  PubMed  Google Scholar 

  167. Yang, C. H., Lee, K. H., Ho, W. K. & Lee, S. H. Inter-spike mitochondrial Ca2+ release enhances high frequency synaptic transmission. J. Physiol. 599, 1567–1594 (2021).

    CAS  PubMed  Google Scholar 

  168. Ma, H., Cai, Q., Lu, W., Sheng, Z. H. & Mochida, S. KIF5B motor adaptor syntabulin maintains synaptic transmission in sympathetic neurons. J. Neurosci. 29, 13019–13029 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  169. Guo, X. et al. The GTPase dMiro is required for axonal transport of mitochondria to Drosophila synapses. Neuron 47, 379–393 (2005).

    CAS  PubMed  Google Scholar 

  170. Schapira, A. H. et al. Mitochondrial complex I deficiency in Parkinson’s disease. Lancet 1, 1269 (1989).

    CAS  PubMed  Google Scholar 

  171. Hsieh, C. H. et al. Functional impairment in Miro degradation and mitophagy is a shared feature in familial and sporadic Parkinson’s disease. Cell Stem Cell 19, 709–724 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  172. Kitada, T. et al. Mutations in the parkin gene cause autosomal recessive juvenile parkinsonism. Nature 392, 605–608 (1998).

    CAS  PubMed  Google Scholar 

  173. Valente, E. M. et al. Hereditary early-onset Parkinson’s disease caused by mutations in PINK1. Science 304, 1158–1160 (2004).

    CAS  PubMed  Google Scholar 

  174. Almikhlafi, M. A. et al. Deletion of DJ-1 in rats affects protein abundance and mitochondrial function at the synapse. Sci. Rep. 10, 13719 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  175. Stauch, K. L. et al. Loss of Pink1 modulates synaptic mitochondrial bioenergetics in the rat striatum prior to motor symptoms: concomitant complex I respiratory defects and increased complex II-mediated respiration. Proteom. Clin. Appl. 10, 1205–1217 (2016).

    CAS  Google Scholar 

  176. Kitada, T. et al. Impaired dopamine release and synaptic plasticity in the striatum of PINK1-deficient mice. Proc. Natl Acad. Sci. USA 104, 11441–11446 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  177. Morais, V. A. et al. Parkinson’s disease mutations in PINK1 result in decreased Complex I activity and deficient synaptic function. EMBO Mol. Med. 1, 99–111 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. Liu, H. F. et al. Combined LRRK2 mutation, aging and chronic low dose oral rotenone as a model of Parkinson’s disease. Sci. Rep. 7, 40887 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  179. Cole, N. B., Dieuliis, D., Leo, P., Mitchell, D. C. & Nussbaum, R. L. Mitochondrial translocation of α-synuclein is promoted by intracellular acidification. Exp. Cell Res. 314, 2076–2089 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  180. Hsu, L. J. et al. α-Synuclein promotes mitochondrial deficit and oxidative stress. Am. J. Pathol. 157, 401–410 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  181. Kramer, M. L. & Schulz-Schaeffer, W. J. Presynaptic α-synuclein aggregates, not Lewy bodies, cause neurodegeneration in dementia with Lewy bodies. J. Neurosci. 27, 1405–1410 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Martin, L. J. et al. Parkinson’s disease α-synuclein transgenic mice develop neuronal mitochondrial degeneration and cell death. J. Neurosci. 26, 41–50 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Nemani, V. M. et al. Increased expression of α-synuclein reduces neurotransmitter release by inhibiting synaptic vesicle reclustering after endocytosis. Neuron 65, 66–79 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  184. Prots, I. et al. α-Synuclein oligomers induce early axonal dysfunction in human iPSC-based models of synucleinopathies. Proc. Natl Acad. Sci. USA 115, 7813–7818 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  185. Reeve, A. K. et al. Mitochondrial dysfunction within the synapses of substantia nigra neurons in Parkinson’s disease. NPJ Parkinsons Dis. 4, 9 (2018).

    PubMed  PubMed Central  Google Scholar 

  186. Ravera, S. et al. Characterization of the mitochondrial aerobic metabolism in the pre- and perisynaptic districts of the SOD1G93A mouse model of amyotrophic lateral sclerosis. Mol. Neurobiol. 55, 9220–9233 (2018).

    CAS  PubMed  Google Scholar 

  187. Bannwarth, S. et al. A mitochondrial origin for frontotemporal dementia and amyotrophic lateral sclerosis through CHCHD10 involvement. Brain 137, 2329–2345 (2014).

    PubMed  PubMed Central  Google Scholar 

  188. Harjuhaahto, S. et al. ALS and Parkinson’s disease genes CHCHD10 and CHCHD2 modify synaptic transcriptomes in human iPSC-derived motor neurons. Neurobiol. Dis. 141, 104940 (2020).

    CAS  PubMed  Google Scholar 

  189. Manzo, E. et al. Glycolysis upregulation is neuroprotective as a compensatory mechanism in ALS. eLife 8, e45114 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  190. Sasaki, S. & Iwata, M. Ultrastructural study of synapses in the anterior horn neurons of patients with amyotrophic lateral sclerosis. Neurosci. Lett. 204, 53–56 (1996).

    CAS  PubMed  Google Scholar 

  191. Sasaki, S. & Iwata, M. Ultrastructural change of synapses of Betz cells in patients with amyotrophic lateral sclerosis. Neurosci. Lett. 268, 29–32 (1999).

    CAS  PubMed  Google Scholar 

  192. Pickett, E. K. et al. Region-specific depletion of synaptic mitochondria in the brains of patients with Alzheimer’s disease. Acta Neuropathol. 136, 747–757 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  193. Hesse, R. et al. Comparative profiling of the synaptic proteome from Alzheimer’s disease patients with focus on the APOE genotype. Acta Neuropathol. Commun. 7, 214 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  194. Beck, S. J. et al. Deregulation of mitochondrial F1FO-ATP synthase via OSCP in Alzheimer’s disease. Nat. Commun. 7, 11483 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  195. Demarest, T. G. et al. Biological sex and DNA repair deficiency drive Alzheimer’s disease via systemic metabolic remodeling and brain mitochondrial dysfunction. Acta Neuropathol. 140, 25–47 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  196. Martino Adami, P. V. et al. Synaptosomal bioenergetic defects are associated with cognitive impairment in a transgenic rat model of early Alzheimer’s disease. J. Cereb. Blood Flow. Metab. 37, 69–84 (2017).

    PubMed  Google Scholar 

  197. Monteiro-Cardoso, V. F. et al. Cardiolipin profile changes are associated to the early synaptic mitochondrial dysfunction in Alzheimer’s disease. J. Alzheimers Dis. 43, 1375–1392 (2015).

    CAS  PubMed  Google Scholar 

  198. Varghese, M. et al. Mitochondrial bioenergetics is defective in presymptomatic Tg2576 AD mice. Transl. Neurosci. 2, https://doi.org/10.2478/s13380-011-0011-8 (2011).

  199. Du, H. et al. Early deficits in synaptic mitochondria in an Alzheimer’s disease mouse model. Proc. Natl Acad. Sci. USA 107, 18670–18675 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  200. Iijima-Ando, K. et al. Mitochondrial mislocalization underlies Aβ42-induced neuronal dysfunction in a Drosophila model of Alzheimer’s disease. PLoS ONE 4, e8310 (2009).

    PubMed  PubMed Central  Google Scholar 

  201. Mhatre, S. D. et al. Synaptic abnormalities in a Drosophila model of Alzheimer’s disease. Dis. Model. Mech. 7, 373–385 (2014).

    PubMed  PubMed Central  Google Scholar 

  202. Quintanilla, R. A., Tapia-Monsalves, C., Vergara, E. H., Pérez, M. J. & Aranguiz, A. Truncated tau induces mitochondrial transport failure through the impairment of TRAK2 protein and bioenergetics decline in neuronal cells. Front. Cell Neurosci. 14, 175 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  203. Zhao, X. L. et al. Expression of β-amyloid induced age-dependent presynaptic and axonal changes in Drosophila. J. Neurosci. 30, 1512–1522 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  204. Han, S., Jeong, Y. Y., Sheshadri, P., Su, X. & Cai, Q. Mitophagy regulates integrity of mitochondria at synapses and is critical for synaptic maintenance. EMBO Rep. 21, e49801 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  205. Fernandez, A. et al. Mitochondrial dysfunction leads to cortical under-connectivity and cognitive impairment. Neuron 102, 1127–1142.e3 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  206. Gokhale, A. et al. Systems analysis of the 22q11.2 microdeletion syndrome converges on a mitochondrial interactome necessary for synapse function and behavior. J. Neurosci. 39, 3561–3581 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  207. Norkett, R., Lesept, F. & Kittler, J. T. DISC1 regulates mitochondrial trafficking in a Miro1-GTP-dependent manner. Front. Cell Dev. Biol. 8, 449 (2020).

    PubMed  PubMed Central  Google Scholar 

  208. Roberts, R. C., Barksdale, K. A., Roche, J. K. & Lahti, A. C. Decreased synaptic and mitochondrial density in the postmortem anterior cingulate cortex in schizophrenia. Schizophr. Res. 168, 543–553 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  209. Huang, N. et al. Reprogramming an energetic AKT-PAK5 axis boosts axon energy supply and facilitates neuron survival and regeneration after injury and ischemia. Curr. Biol. 31, 3098–3114.e7 (2021).

    CAS  PubMed  Google Scholar 

  210. Chamberlain, K. A. et al. Oligodendrocytes enhance axonal energy metabolism by deacetylation of mitochondrial proteins through transcellular delivery of SIRT2. Neuron https://doi.org/10.1016/j.neuron.2021.08.011 (2021).

    Article  PubMed  Google Scholar 

  211. Balietti, M. et al. Early selective vulnerability of synapses and synaptic mitochondria in the hippocampal CA1 region of the Tg2576 mouse model of Alzheimer’s disease. J. Alzheimers Dis. 34, 887–896 (2013).

    CAS  PubMed  Google Scholar 

  212. Trushina, E. et al. Defects in mitochondrial dynamics and metabolomic signatures of evolving energetic stress in mouse models of familial Alzheimer’s disease. PLoS ONE 7, e32737 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  213. Györffy, B. A. et al. Synaptic mitochondrial dysfunction and septin accumulation are linked to complement-mediated synapse loss in an Alzheimer’s disease animal model. Cell Mol. Life Sci. 77, 5243–5258 (2020).

    PubMed  PubMed Central  Google Scholar 

  214. Völgyi, K. et al. Mitochondrial proteome changes correlating with β-amyloid accumulation. Mol. Neurobiol. 54, 2060–2078 (2017).

    PubMed  Google Scholar 

  215. Dragicevic, N. et al. Mitochondrial amyloid-β levels are associated with the extent of mitochondrial dysfunction in different brain regions and the degree of cognitive impairment in Alzheimer’s transgenic mice. J. Alzheimers Dis. 20, S535–S550 (2010).

    PubMed  Google Scholar 

  216. Espino de la Fuente-Muñoz, C. et al. Age-dependent decline in synaptic mitochondrial function is exacerbated in vulnerable brain regions of female 3xTg-AD mice. Int. J. Mol. Sci. 21, 8727 (2020).

    PubMed Central  Google Scholar 

  217. Machamer, J. B., Woolums, B. M., Fuller, G. G. & Lloyd, T. E. FUS causes synaptic hyperexcitability in Drosophila dendritic arborization neurons. Brain Res. 1693, 55–66 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  218. Virlogeux, A. et al. Reconstituting corticostriatal network on-a-chip reveals the contribution of the presynaptic compartment to Huntington’s disease. Cell Rep. 22, 110–122 (2018).

    CAS  PubMed  Google Scholar 

  219. Yano, H. et al. Inhibition of mitochondrial protein import by mutant huntingtin. Nat. Neurosci. 17, 822–831 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  220. Baranov, S. V. et al. Mitochondria modulate programmed neuritic retraction. Proc. Natl Acad. Sci. USA 116, 650–659 (2019).

    CAS  PubMed  Google Scholar 

  221. Orr, A. L. et al. N-terminal mutant huntingtin associates with mitochondria and impairs mitochondrial trafficking. J. Neurosci. 28, 2783–2792 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  222. Ferreirinha, F. et al. Axonal degeneration in paraplegin-deficient mice is associated with abnormal mitochondria and impairment of axonal transport. J. Clin. Invest. 113, 231–242 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  223. Gorski, K. et al. Quantitative changes in the mitochondrial proteome of cerebellar synaptosomes from preclinical cystatin B-deficient mice. Front. Mol. Neurosci. 13, 570640 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  224. Ebrahimi-Fakhari, D. et al. Impaired mitochondrial dynamics and mitophagy in neuronal models of tuberous sclerosis complex. Cell Rep. 17, 1053–1070 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  225. Jiang, S. et al. Mfn2 ablation causes an oxidative stress response and eventual neuronal death in the hippocampus and cortex. Mol. Neurodegener. 13, 5 (2018).

    PubMed  PubMed Central  Google Scholar 

  226. Trevisan, T. et al. Manipulation of mitochondria dynamics reveals separate roles for form and function in mitochondria distribution. Cell Rep. 23, 1742–1753 (2018).

    CAS  PubMed  Google Scholar 

  227. Brickley, K. & Stephenson, F. A. Trafficking kinesin protein (TRAK)-mediated transport of mitochondria in axons of hippocampal neurons. J. Biol. Chem. 286, 18079–18092 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  228. Glater, E. E., Megeath, L. J., Stowers, R. S. & Schwarz, T. L. Axonal transport of mitochondria requires milton to recruit kinesin heavy chain and is light chain independent. J. Cell Biol. 173, 545–557 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  229. Stowers, R. S., Megeath, L. J., Górska-Andrzejak, J., Meinertzhagen, I. A. & Schwarz, T. L. Axonal transport of mitochondria to synapses depends on milton, a novel Drosophila protein. Neuron 36, 1063–1077 (2002).

    CAS  PubMed  Google Scholar 

  230. Morciano, G. et al. Use of luciferase probes to measure ATP in living cells and animals. Nat. Protoc. 12, 1542–1562 (2017).

    CAS  PubMed  Google Scholar 

  231. Berg, J., Hung, Y. P. & Yellen, G. A genetically encoded fluorescent reporter of ATP:ADP ratio. Nat. Methods 6, 161–166 (2009). This study reports generation of the fluorescent energy sensor Perceval for assessing intracellular ATP/ADP ratios in live cells.

    CAS  PubMed  PubMed Central  Google Scholar 

  232. Tantama, M., Martínez-François, J. R., Mongeon, R. & Yellen, G. Imaging energy status in live cells with a fluorescent biosensor of the intracellular ATP-to-ADP ratio. Nat. Commun. 4, 2550 (2013). This study optimizes the genetically encoded fluorescent biosensor Perceval into a high-range version called ‘PercevalHR’ that can sense intracellular ATP/ADP ratios.

    PubMed  Google Scholar 

  233. Lobas, M. A. et al. A genetically encoded single-wavelength sensor for imaging cytosolic and cell surface ATP. Nat. Commun. 10, 711 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  234. Nakano, M., Imamura, H., Nagai, T. & Noji, H. Ca2+ regulation of mitochondrial ATP synthesis visualized at the single cell level. ACS Chem. Biol. 6, 709–715 (2011).

    CAS  PubMed  Google Scholar 

  235. Obashi, K. & Okabe, S. Regulation of mitochondrial dynamics and distribution by synapse position and neuronal activity in the axon. Eur. J. Neurosci. 38, 2350–2363 (2013).

    PubMed  Google Scholar 

  236. Tanaka, Y. et al. Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria. Cell 93, 1147–1158 (1998).

    CAS  PubMed  Google Scholar 

  237. Pilling, A. D., Horiuchi, D., Lively, C. M. & Saxton, W. M. Kinesin-1 and dynein are the primary motors for fast transport of mitochondria in Drosophila motor axons. Mol. Biol. Cell 17, 2057–2068 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  238. Zhao, Y. et al. Metaxins are core components of mitochondrial transport adaptor complexes. Nat. Commun. 12, 83 (2021).

    CAS  PubMed  PubMed Central  Google Scholar 

  239. Frederick, R. L., McCaffery, J. M., Cunningham, K. W., Okamoto, K. & Shaw, J. M. Yeast Miro GTPase, Gem1p, regulates mitochondrial morphology via a novel pathway. J. Cell Biol. 167, 87–98 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  240. Birsa, N., Norkett, R., Higgs, N., Lopez-Domenech, G. & Kittler, J. T. Mitochondrial trafficking in neurons and the role of the Miro family of GTPase proteins. Biochem. Soc. Trans. 41, 1525–1531 (2013).

    CAS  PubMed  Google Scholar 

  241. Cai, Q., Gerwin, C. & Sheng, Z. H. Syntabulin-mediated anterograde transport of mitochondria along neuronal processes. J. Cell Biol. 170, 959–969 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  242. Russo, G. J. et al. Drosophila Miro is required for both anterograde and retrograde axonal mitochondrial transport. J. Neurosci. 29, 5443–5455 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  243. van Spronsen, M. et al. TRAK/Milton motor-adaptor proteins steer mitochondrial trafficking to axons and dendrites. Neuron 77, 485–502 (2013).

    PubMed  Google Scholar 

  244. Drerup, C. M., Herbert, A. L., Monk, K. R. & Nechiporuk, A. V. Regulation of mitochondria-dynactin interaction and mitochondrial retrograde transport in axons. eLife 6, e22234 (2017).

    PubMed  PubMed Central  Google Scholar 

  245. van Bergeijk, P., Adrian, M., Hoogenraad, C. C. & Kapitein, L. C. Optogenetic control of organelle transport and positioning. Nature 518, 111–114 (2015). This optogenetic study shows that selectively recruiting KIF5 or SNPH to axonal mitochondria shifts the balance between motile and stationary pools.

    PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

The authors thank the laboratories and scientists who contributed to data and discoveries discussed here, and J. C. Roney for critical reading and editing. The authors apologize to those colleagues whose work could not be cited because of space limitations. This work was supported by the Intramural Research Program of NINDS, NIH ZIA NS003029 and ZIA NS002946 (Z.-H.S.).

Author information

Authors and Affiliations

Authors

Contributions

The authors contributed equally to all aspects of the article.

Corresponding author

Correspondence to Zu-Hang Sheng.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information

Nature Reviews Neuroscience thanks D. Chan, J. Kittler, W. Kunz and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Publisher’s note

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

Glossary

Action potential

A brief reversal of electrical polarization across the neuronal membrane that results from the opening and closing of voltage-gated ion channels and produces the nerve impulse that permits communication between neurons.

Glycolysis

A partial glucose metabolism that occurs in the cytosol or on membrane surfaces. Glycolysis converts a six-carbon glucose into two three-carbon pyruvates, with a net gain of two adenosine triphosphate (ATP) molecules and two NADH molecules. Pyruvate enters the tricarboxylic acid cycle in the mitochondrial matrix or is reduced to lactate.

Oxidative phosphorylation

Highly efficient energetic metabolism of pyruvate that occurs in the inner mitochondrial membrane. Oxidative phosphorylation generates adenosine triphosphate (ATP) through the electron transport chain and the actions of ATP synthase.

Synaptic vesicle recycling

A synaptic vesicle cycle that occurs at presynaptic terminals, encompassing synaptic vesicle endocytosis, refilling the vesicle with neurotransmitters, re-formation of synaptic vesicle pools, mobilization to release sites and the release of neurotransmitters via exocytosis.

Synaptoenergetics

The status of the bioenergetics processes involved in the generation and consumption of adenosine triphosphate (ATP) at the synapse. Synaptoenergetics is maintained and adapted through activity-dependent and/or energy-sensitive regulation of local energy metabolism to sustain synaptic efficacy and plasticity.

Electron transport chain

A series of protein complexes residing on the mitochondrial inner membrane that transfer electrons through electron carriers to form a proton gradient that drives the creation of adenosine triphosphate (ATP).

Tricarboxylic acid (TCA) cycle

A series of enzymatic reactions that occur in a closed loop in the mitochondrial matrix. The TCA cycle completely breaks down glucose and oxidizes acetyl coenzyme A to generate the NADH and FADH2 that are required for the electron transport chain.

Glycolytic metabolon

A transient protein complex of sequential glycolytic enzymes that enables metabolites to channel directly from one enzyme to the next to catalyse stepwise glycolytic reactions.

Energy stress

An imbalanced energetic status caused by insufficient adenosine triphosphate (ATP) supply and/or enhanced energy consumption, resulting in an increased adenosine diphosphate (ADP) to ATP ratio.

Glucose transporter

One of a group of membrane proteins that take glucose from extracellular spaces into cells.

AMP-activated protein kinase

(AMPK). A master cellular energy stress sensor that is activated when intracellular adenosine triphosphate (ATP) becomes depleted or ATP consumption is elevated.

Mitochondrial fission and fusion

Mitochondrial membrane dynamic with the ability to join two mitochondria together (fusion) or separate one mitochondrion into two mitochondria (fission) mediated by multiple fusion or fission proteins. Fission and fusion dynamics control mitochondrial morphology and size and maintain mitochondrial integrity, distribution and metabolism.

Mitophagy

The selective elimination of damaged mitochondria by the autophagolysosomal system.

Long-term potentiation

(LTP). A persistent type of synaptic plasticity that induces a long-lasting increase in the efficacy of synaptic transmission.

Active zones

Specialized synaptic vesicle recycling regions that display an electron-dense thickening of the presynaptic membrane, together with a cluster of synaptic vesicles, in an area that apposes the postsynaptic region.

Crista

An invaginated structure of the mitochondrial inner membrane that extends deep into the matrix to increase the functional surface area of the inner membrane for cellular respiration.

Motor proteins

A class of molecular motors that drive intracellular trafficking of organelles along cytoskeletal filaments by the hydrolysis of adenosine triphosphate (ATP).

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Li, S., Sheng, ZH. Energy matters: presynaptic metabolism and the maintenance of synaptic transmission. Nat Rev Neurosci 23, 4–22 (2022). https://doi.org/10.1038/s41583-021-00535-8

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41583-021-00535-8

This article is cited by

Search

Quick links

Nature Briefing

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

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