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Mitochondrial transport in neurons: impact on synaptic homeostasis and neurodegeneration

Key Points

  • Mitochondria are essential for neuronal function and survival. Mitochondria are commonly found in synaptic terminals, where they help to maintain neurotransmission by producing ATP and buffering Ca2+.

  • Mitochondrial transport and distribution in neurons is efficiently regulated in response to changes in neuronal activity and various physiological and pathological states.

  • Neuronal mitochondria undergo dynamic and bidirectional transport along neuronal processes, frequently changing direction, pausing or switching to persistent docking.

  • These complex mitochondrial mobility patterns are a result of mitochondrial coupling to anterograde kinesin motors of the KIF5 family and to the retrograde motor dynein, as well as to docking and anchoring machineries, including syntaphilin.

  • Mitochondria attach to the motors by associating with their respective motor adaptor proteins and mitochondrial receptors. These motor–adaptor–receptor complexes ensure targeted trafficking of mitochondria and precise regulation of their mobility.

  • The KIF5–Milton–MIRO complex constitutes a mitochondrial transport machinery, through which a MIRO Ca2+-sensing pathway mediates the suppression of mitochondrial mobility in response to increased action potential firing rates or the activation of glutamate receptors.

  • Syntaphilin acts as a 'static anchor' for axonal mitochondria. Deleting the syntaphilin gene resulted in a robust increase in the percentage of mobile axonal mitochondria relative to wild-type neurons.

  • Elaborate mitochondrial quality-control systems maintain mitochondrial integrity and function. These include transport, fusion, fission and turnover via mitophagy and constitute an interdependent system of mitochondrial dynamics.

  • It is well documented that mitochondrial dysfunction, changes in mitochondrial dynamics and mobility, and perturbation of mitochondrial turnover are involved in the pathology of some major neurodegenerative and neurological disorders.

  • Identification of the molecules involved in linking mitochondrial transport, fusion, fission and mitophagy will advance our understanding of the cellular mechanisms that regulate mitochondrial quality control and thus human neurodegenerative diseases.

Abstract

Mitochondria have a number of essential roles in neuronal function. Their complex mobility patterns within neurons are characterized by frequent changes in direction. Mobile mitochondria can become stationary or pause in regions that have a high metabolic demand and can move again rapidly in response to physiological changes. Defects in mitochondrial transport are implicated in the pathogenesis of several major neurological disorders. Research into the mechanisms that regulate mitochondrial transport is thus an important emerging frontier.

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Figure 1: Mitochondrial transport in neurons.
Figure 2: KIF5-driven mitochondrial transport.
Figure 3: Mitochondrial docking and synaptic homeostasis.

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References

  1. Nicholls, D. G. & Budd, S. L. Mitochondria and neuronal survival. Physiol. Rev. 80, 315–360 (2000).

    Article  CAS  PubMed  Google Scholar 

  2. Verstreken, P. et al. Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions. Neuron 47, 365–378 (2005). This paper shows that in D. melanogaster with mutant DRP1, the loss of mitochondria from neuromuscular junctions results in faster depletion of synaptic vesicles during prolonged pulse train stimulation owing to a specific defect in mobilizing reserve pool vesicles.

    Article  CAS  PubMed  Google Scholar 

  3. Lee, C. W. & Peng, H. B. The function of mitochondria in presynaptic development at the neuromuscular junction. Mol. Biol. Cell 19, 150–158 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. 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).

    Article  CAS  PubMed  Google Scholar 

  5. Werth, J. L. & Thayer, S. A. Mitochondria buffer physiological calcium loads in cultured rat dorsal root ganglion neurons. J. Neurosci. 14, 348–356 (1994).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. Medler, K. & Gleason, E. L. Mitochondrial Ca2+ buffering regulates synaptic transmission between retinal amacrine cells. J. Neurophysiol. 87, 1426–1439 (2002).

    Article  CAS  PubMed  Google Scholar 

  9. David, G. & Barrett, E. F. Mitochondrial Ca2+ uptake prevents desynchronization of quantal release and minimizes depletion during repetitive stimulation of mouse motor nerve terminals. J. Physiol. 548, 425–438 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  10. Talbot, J. D., David, G. & Barrett, E. F. Inhibition of mitochondrial Ca2+ uptake affects phasic release from motor terminals differently depending on external [Ca2+]. J. Neurophysiol. 90, 491–502 (2003).

    Article  CAS  PubMed  Google Scholar 

  11. Levy, M., Faas, G. C., Saggau, P., Craigen, W. J. & Sweatt, J. D. Mitochondrial regulation of synaptic plasticity in the hippocampus. J. Biol. Chem. 278, 17727–17734 (2003).

    Article  CAS  PubMed  Google Scholar 

  12. Kang, J. S. et al. Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation. Cell 132, 137–148 (2008). By using genetic mouse models and time-lapse imaging this study identifies syntaphilin as a 'static anchor' for axonal mitochondria.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Hollenbeck, P. J. & Saxton, W. M. The axonal transport of mitochondria. J. Cell Sci. 118, 5411–5419 (2005).

    Article  CAS  PubMed  Google Scholar 

  14. Bogan, N. & Cabot, J. B. Light and electron microscopic analyses of intraspinal axon collaterals of sympathetic preganglionic neurons. Brain Res. 541, 241–251 (1991).

    Article  CAS  PubMed  Google Scholar 

  15. Fabricius, C., Berthold, C. H. & Rydmark, M. Axoplasmic organelles at nodes of Ranvier. II. Occurrence and distribution in large myelinated spinal cord axons of the adult cat. J. Neurocytol. 22, 941–954 (1993).

    Article  CAS  PubMed  Google Scholar 

  16. 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 

  17. Mutsaers, S. E. & Carroll, W. M. Focal accumulation of intra-axonal mitochondria in demyelination of the cat optic nerve. Acta Neuropathol. 96, 139–143 (1998).

    Article  CAS  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Li, Z., Okamoto, K., Hayashi, Y. & Sheng, M. The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses. Cell 119, 873–887 (2004).

    Article  CAS  PubMed  Google Scholar 

  20. Zhang, C. L., Ho, P. L., Kintner, D. B., Sun, D. & Chiu, S. Y. Activity-dependent regulation of mitochondrial motility by calcium and Na/K-ATPase at nodes of Ranvier of myelinated nerves. J. Neurosci. 30, 3555–3566 (2010). This study demonstrates a highly localized elevation of axonal Ca2+ levels and reduced mitochondrial mobility at individual nodes of Ranvier during a brief train of action potentials.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. 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). This study shows that mobile mitochondria are recruited to stationary pools in response to acute application of glutamate to cultured neurons. Mitochondria are also changed from an elongated to a rounded morphology.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Macaskill, A. F. et al. Miro1 is a calcium sensor for glutamate receptor-dependent localization of mitochondria at synapses. Neuron 61, 541–555 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Saotome, M. et al. Bidirectional Ca2+-dependent control of mitochondrial dynamics by the Miro GTPase. Proc. Natl Acad. Sci. USA 105, 20728–20733 (2008). References 23, 24 and 25 independently identified MIRO as a Ca2+ sensor, providing a mechanism for the underlying Ca2+-dependent regulation of mitochondrial mobility.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Yi, M., Weaver, D. & Hajnóczky, G. Control of mitochondrial motility and distribution by the calcium signal: a homeostatic circuit. J. Cell Biol. 167, 661–672 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  28. Chan, D. C. Mitochondria: dynamic organelles in disease, aging, and development. Cell 125, 1241–1252 (2006).

    Article  CAS  PubMed  Google Scholar 

  29. Stokin, G. B. & Goldstein, L. S. Axonal transport and Alzheimer's disease. Annu. Rev. Biochem. 75, 607–627 (2006).

    Article  CAS  PubMed  Google Scholar 

  30. Schon, E. A. & Przedborski, S. Mitochondria: the next (neurode) generation. Neuron 70, 1033–1053 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Miller, K. E. & Sheetz, M. P. Direct evidence for coherent low velocity axonal transport of mitochondria. J. Cell Biol. 173, 373–381 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Misgeld, T., Kerschensteiner, M., Bareyre, F. M., Burgess, R. W. & Lichtman, J. W. Imaging axonal transport of mitochondria in vivo. Nature Methods 4, 559–561 (2007). This study develops an elegant tool to visualize axonal mitochondrial transport in living mice and explanted nervous tissue.

    Article  CAS  PubMed  Google Scholar 

  33. Hirokawa, N., Niwa, S. & Tanaka, Y. Molecular motors in neurons: transport mechanisms and roles in brain function, development, and disease. Neuron 68, 610–638 (2010).

    Article  CAS  PubMed  Google Scholar 

  34. Martin, M. et al. Cytoplasmic dynein, the dynactin complex, and kinesin are interdependent and essential for fast axonal transport. Mol. Biol. Cell 10, 3717–3728 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. 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). This is the first genetic mouse study showing that KIF5 motors are essential for mitochondrial transport.

    Article  CAS  PubMed  Google Scholar 

  36. Górska-Andrzejak, J. et al. Mitochondria are redistributed in Drosophila photoreceptors lacking milton, a kinesin-associated protein. J. Comp. Neurol. 463, 372–388 (2003).

    Article  CAS  PubMed  Google Scholar 

  37. 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). This study provides the genetic evidence in the D. melanogaster nervous system that dynein motors have a crucial role in mitochondrial retrograde transport in axons.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  38. MacAskill, A. F. & Kittler, J. T. Control of mitochondrial transport and localization in neurons. Trends Cell Biol. 20, 102–112 (2010).

    Article  CAS  PubMed  Google Scholar 

  39. Kanai, Y. et al. KIF5C, a novel neuronal kinesin enriched in motor neurons. J. Neurosci. 20, 6374–6384 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Hirokawa, N. et al. Kinesin associates with anterogradely transported membranous organelles in vivo. J. Cell Biol. 114, 295–302 (1991). This study shows that KIF5 motors are attached to brain mitochondria.

    Article  CAS  PubMed  Google Scholar 

  41. Cai, Q., Gerwin, C. & Sheng, Z.-H. Syntabulin-mediated anterograde transport of mitochondria along neuronal processes. J. Cell Biol. 170, 959–969 (2005). This study identifies syntabulin as a second prominent KIF5 motor adaptor for mitochondria. Syntabulin loss-of-function or interference of the syntaphilin–KIF5 interaction reduces anterograde, but not retrograde, mitochondrial transport along axons.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Nangaku, M. et al. KIF1B, a novel microtubule plus end-directed monomeric motor protein for transport of mitochondria. Cell 79, 1209–1220 (1994).

    Article  CAS  PubMed  Google Scholar 

  43. Tanaka, K., Sugiura, Y., Ichishita, R., Mihara, K. & Oka, T. KLP6: a newly identified kinesin that regulates the morphology and transport of mitochondria in neuronal cells. J. Cell Sci. 124, 2457–2465 (2011).

    Article  CAS  PubMed  Google Scholar 

  44. 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). This paper reports that D. melanogaster photoreceptors that express mutant Milton show aberrant synaptic transmission owing to a reduced distribution of mitochondria at synapses.

    Article  CAS  PubMed  Google Scholar 

  45. Fransson, A., Ruusala, A. & Aspenström, P. Atypical Rho GTPases have roles in mitochondrial homeostasis and apoptosis. J. Biol. Chem. 278, 6495–6502 (2003).

    Article  CAS  PubMed  Google Scholar 

  46. Fransson, S., Ruusala, A. & Aspenström, P. The atypical Rho GTPases Miro-1 and Miro-2 have essential roles in mitochondrial trafficking. Biochem. Biophys. Res. Commun. 344, 500–510 (2006).

    Article  CAS  PubMed  Google Scholar 

  47. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  48. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  49. Guo, X. et al. The GTPase dMiro is required for axonal transport of mitochondria to Drosophila synapses. Neuron 47, 379–393 (2005). In this paper, the authors show that a MIRO mutant results in the chronic loss of mitochondria from neuromuscular junctions, a reduction in Ca2+ buffering capacity and impaired neurotransmitter release during prolonged stimulation.

    Article  CAS  PubMed  Google Scholar 

  50. Brickley, K., Smith, M. J., Beck, M. & Stephenson, F. A. GRIF-1 and OIP106, members of a novel gene family of coiled-coil domain proteins: association in vivo and in vitro with kinesin. J. Biol. Chem. 280, 14723–14732 (2005).

    Article  CAS  PubMed  Google Scholar 

  51. Smith, M. J., Pozo, K., Brickley, K. & Stephenson, F. A. Mapping the GRIF-1 binding domain of the kinesin, KIF5C, substantiates a role for GRIF-1 as an adaptor protein in the anterograde trafficking of cargoes. J. Biol. Chem. 281, 27216–27228 (2006).

    Article  CAS  PubMed  Google Scholar 

  52. Grishin, A., Li, H., Levitan, E. S. & Zaks-Makhina, E. Identification of γ-aminobutyric acid receptor-interacting factor 1 (TRAK2) as a trafficking factor for the K+ channel Kir2.1. J. Biol. Chem. 281, 30104–30111 (2006).

    Article  CAS  PubMed  Google Scholar 

  53. Kirk, E., Chin, L. S. & Li, L. GRIF1 binds Hrs and is a new regulator of endosomal trafficking. J. Cell Sci. 119, 4689–4701 (2006).

    Article  CAS  PubMed  Google Scholar 

  54. Webber, E., Li, L. & Chin, L. S. Hypertonia-associated protein Trak1 is a novel regulator of endosome-to-lysosome trafficking. J. Mol. Biol. 382, 638–651 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  55. MacAskill, A. F., Brickley, K., Stephenson, F. A. & Kittler, J. T. GTPase dependent recruitment of Grif-1by Miro1 regulates mitochondrial trafficking in hippocampal neurons. Mol. Cell. Neurosci. 40, 301–312 (2009).

    Article  CAS  PubMed  Google Scholar 

  56. 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). References 55 and 56 provide evidence that elevating MIRO1 levels enhances the recruitment of the TRAK2–KIF5 transport complex to mitochondria and that knocking down TRAK1 or expressing dominant-negative TRAK1 mutants results in impaired mitochondrial mobility in axons.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  57. Ikuta, J. et al. Fasciculation and elongation protein zeta-1 (FEZ1) participates in the polarization of hippocampal neuron by controlling the mitochondrial motility. Biochem. Biophys. Res. Commun. 353, 127–132 (2007).

    Article  CAS  PubMed  Google Scholar 

  58. Fujita T. et al. Axonal guidance protein FEZ1 associates with tubulin and kinesin motor protein to transport mitochondria in neurites of NGF-stimulated PC12 cells. Biochem. Biophys. Res. Commun. 361, 605–610 (2007).

    Article  CAS  PubMed  Google Scholar 

  59. Cho, K. I. et al. Association of the kinesin-binding domain of RanBP2 to KIF5B and KIF5C determines mitochondria localization and function. Traffic 8, 1722–1735 (2007).

    Article  CAS  PubMed  Google Scholar 

  60. Schwarzer, C., Barnikol-Watanabe, S., Thinnes, F. P. & Hilschmann, N. Voltage-dependent anion-selective channel (VDAC) interacts with the dynein light chain Tctex1 and the heat-shock protein PBP74. Int. J. Biochem. Cell Biol. 34, 1059–1070 (2002).

    Article  CAS  PubMed  Google Scholar 

  61. King, S. J. & Schroer, T. A. Dynactin increases the processivity of the cytoplasmic dynein motor. Nature Cell Biol. 2, 20–24 (2000).

    Article  CAS  PubMed  Google Scholar 

  62. Haghnia, M. et al. Dynactin is required for coordinated bidirectional motility, but not for dynein membrane attachment. Mol. Biol. Cell 18, 2081–2089 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. Cai, Q. et al. Snapin-regulated late endosomal transport is critical for efficient autophagy-lysosomal function in neurons. Neuron 68, 73–86 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Russo, G. J. et al. Drosophila Miro is required for both anterograde and retrograde axonal mitochondrial transport. J. Neurosci. 29, 5443–5455 (2009). This study suggests that MIRO promotes either kinesin- or dynein-mediated movement during a neuronal signal that dictates the net transport direction of mitochondria.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  65. Hirokawa, N., Sato-Yoshitake, R., Yoshida, T. & Kawashima, T. Brain dynein (MAP1C) localizes on both anterogradely and retrogradely transported membranous organelles in vivo. J. Cell Biol. 111, 1027–1037 (1990).

    Article  CAS  PubMed  Google Scholar 

  66. Ligon, L. A., Tokito, M., Finklestein, J. M., Grossman, F. E. & Holzbaur, E. L. A direct interaction between cytoplasmic dynein and kinesin I may coordinate motor activity. J. Biol. Chem. 279, 19201–19208 (2004).

    Article  CAS  PubMed  Google Scholar 

  67. Welte, M. A. Bidirectional transport along microtubules. Curr. Biol. 14, R525–R537 (2004).

    Article  CAS  PubMed  Google Scholar 

  68. Mallik, R., Petrov, D., Lex, S. A., King, S. J. & Gross, S. P. Building complexity: an in vitro study of cytoplasmic dynein with in vivo implications. Curr. Biol. 15, 2075–2085 (2005).

    Article  CAS  PubMed  Google Scholar 

  69. Horiuchi, D., Barkus, R. V., Pilling, A. D., Gassman, A. & Saxton, W. M. APLIP1, a kinesin binding JIP-1/JNK scaffold protein, influences the axonal transport of both vesicles and mitochondria in Drosophila. Curr. Biol. 15, 2137–2141 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  70. Misko, A., Jiang, S., Wegorzewska, I., Milbrandt, J. & Baloh, R. H. Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex. J. Neurosci. 30, 4232–4240 (2010). This study highlights a role of the MIRO2–MFN2 complex in regulating the processivity of kinesin or in coordinating the switch between kinesin and dynein.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  71. 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).

    Article  CAS  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. Naisbitt, S. et al. Interaction of the postsynaptic density-95/guanylate kinase domain-associated protein complex with a light chain of myosin-V and dynein. J. Neurosci. 20, 4524–4534 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  75. 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). References 74 and 75 provide compelling evidence that NGF can regulate mitochondrial mobility by influencing static interactions between mitochondria and actin and that inhibiting actin-based myosin motors results in increased mitochondrial mobility.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Hubley, M. J., Locke, B. R. & Moerland, T. S. The effects of temperature, pH, and magnesium on the diffusion coefficient of ATP in solutions of physiological ionic strength. Biochim. Biophys. Acta 1291, 115–121 (1996).

    Article  PubMed  Google Scholar 

  77. Hirokawa, N. Cross-linker system between neurofilaments, microtubules, and membranous organelles in frog axons revealed by the quick-freeze, deep-etching method. J. Cell Biol. 94, 129–142 (1982). This study provides the first morphological evidence for the crossbridges between axonal mitochondria and microtubules or neurofilaments.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  78. Lindén, M., Nelson, B. D., Loncar, D. & Leterrier, J. F. Studies on the interaction between mitochondria and the cytoskeleton. J. Bioenerg. Biomembr. 21, 507–518 (1989).

    Article  PubMed  Google Scholar 

  79. Jung, D., Filliol, D., Miehe, M. & Rendon, A. Interaction of brain mitochondria with microtubules reconstituted from brain tubulin and MAP2 or TAU. Cell Motil. Cytoskeleton 24, 245–255 (1993).

    Article  CAS  PubMed  Google Scholar 

  80. Price, R. L., Lasek, R. J. & Katz, M. J. Microtubules have special physical associations with smooth endoplasmic reticula and mitochondria in axons. Brain Res. 540, 209–216 (1991).

    Article  CAS  PubMed  Google Scholar 

  81. Chen, Y. M., Gerwin, C. & Sheng, Z.-H. Dynein light chain LC8 regulates syntaphilin-mediated mitochondrial docking in axons. J. Neurosci. 29, 9429–9438 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Sung, J. Y. et al. WAVE1 controls neuronal activity-induced mitochondrial distribution in dendritic spines. Proc. Natl Acad. Sci. USA. 105, 3112–3116 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  83. Wagner, O. I. et al. Mechanisms of mitochondria-neurofilament interactions. J. Neurosci. 23, 9046–9058 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  84. Mironov, S. L. ADP regulates movements of mitochondria in neurons. Biophys. J. 92, 2944–2952 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  85. Chang, D. T., Honick, A. S. & Reynolds, I. J. Mitochondrial trafficking to synapses in cultured primary cortical neurons. J. Neurosci. 26, 7035–7045 (2006). This study demonstrates that mitochondrial mobility is regulated in response to changes in synaptic activity.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  86. Cai, Q. & Sheng, Z. H. Moving or stopping mitochondria: Miro as a traffic cop by sensing calcium. Neuron 61, 493–496 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Chang, K. T., Niescier, R. F. & Min, K. T. Mitochondrial matrix Ca2+ as an intrinsic signal regulating mitochondrial motility in axons. Proc. Natl Acad. Sci. USA 108, 15456–15461 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  88. Han, X. J. et al. CaM kinase I α-induced phosphorylation of Drp1 regulates mitochondrial morphology. J. Cell Biol. 182, 573–585 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Chen, S., Owens, G. C., Crossin, K. L. & Edelman, D. B. Serotonin stimulates mitochondrial transport in hippocampal neurons. Mol. Cell. Neurosci. 36, 472–483 (2007).

    Article  CAS  PubMed  Google Scholar 

  90. Chen, S. et al. Dopamine inhibits mitochondrial motility in hippocampal neurons. PLoS ONE 3, e2804 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  91. 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).

    Article  CAS  PubMed  Google Scholar 

  92. Zanelli, S. A. et al. Nitric oxide impairs mitochondrial movement in cortical neurons during hypoxia. J. Neurochem. 97, 724–736 (2006).

    Article  CAS  PubMed  Google Scholar 

  93. Stamer, K., Vogel, R., Thies, E., Mandelkow, E. & Mandelkow, E. M. Tau blocks traffic of organelles, neurofilaments, and APP vesicles in neurons and enhances oxidative stress. J. Cell Biol. 156, 1051–1063 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Dubey, M., Chaudhury, P., Kabiru, H. & Shea, T. B. Tau inhibits anterograde axonal transport and perturbs stability in growing axonal neurites in part by displacing kinesin cargo: neurofilaments attenuate tau-mediated neurite instability. Cell Motil. Cytoskeleton 65, 89–99 (2008).

    Article  CAS  PubMed  Google Scholar 

  95. Stoothoff, W. et al. Differential effect of three-repeat and four-repeat tau on mitochondrial axonal transport. J. Neurochem. 111, 417–427 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  96. Trinczek, B., Ebneth, A., Mandelkow, E. M. & Mandelkow, E. Tau regulates the attachment/detachment but not the speed of motors in microtubule-dependent transport of single vesicles and organelles. J. Cell Sci. 112, 2355–2367 (1999).

    CAS  PubMed  Google Scholar 

  97. Mandelkow, E. M., Thies, E., Trinczek, B., Biernat, J. & Mandelkow, E. MARK/PAR1 kinase is a regulator of microtubule-dependent transport in axons. J. Cell Biol. 167, 99–110 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  98. Vossel, K. A. et al. Tau reduction prevents Aβ-induced defects in axonal transport. Science 330, 198 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  99. Dixit, R., Ross, J. L., Goldman, Y. E. & Holzbaur, E. L. Differential regulation of dynein and kinesin motor proteins by tau. Science 319, 1086–1089 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  100. Jiménez-Mateos, E. M., González-Billault, C., Dawson, H. N., Vitek, M. P. & Avila, J. Role of MAP1B in axonal retrograde transport of mitochondria. Biochem. J. 397, 53–59 (2006).

    Article  PubMed  PubMed Central  Google Scholar 

  101. Ohno, N. et al. Myelination and axonal electrical activity modulate the distribution and motility of mitochondria at CNS nodes of Ranvier. J. Neurosci. 31, 7249–7258 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  102. Craner, M. J. et al. Molecular changes in neurons in multiple sclerosis: altered axonal expression of Nav1.2 and Nav1.6 sodium channels and Na+/Ca2+ exchanger. Proc. Natl Acad. Sci. USA 101, 8168–8173 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  103. Andrews, H. et al. Increased axonal mitochondrial activity as an adaptation to myelin deficiency in the Shiverer mouse. J. Neurosci. Res. 83, 1533–1539 (2006).

    Article  CAS  PubMed  Google Scholar 

  104. Hogan, V. et al. Increase in mitochondrial density within axons and supporting cells in response to demyelination in the Plp1 mouse model. J. Neurosci. Res. 87, 452–459 (2009).

    Article  CAS  PubMed  Google Scholar 

  105. Kiryu-Seo, S., Ohno, N., Kidd, G. J., Komuro, H. & Trapp, B. D. Demyelination increases axonal stationary mitochondrial size and the speed of axonal mitochondrial transport. J. Neurosci. 30, 6658–6666 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  106. Shepherd, G. M. & Harris, K. M. Three-dimensional structure and composition of CA3-CA1 axons in rat hippocampal slices: implications for presynaptic connectivity and compartmentalization. J. Neurosci. 18, 8300–8310 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  107. Rowland, K. C., Irby, N. K. & Spirou, G. A. Specialized synapse-associated structures within the calyx of Held. J. Neurosci. 20, 9135–9144 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  108. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. Gross, N. J., Getz, G. S. & Rabinowitz, M. Apparent turnover of mitochondrial deoxyribonucleic acid and mitochondrial phospholipids in the tissues of the rat. J. Biol. Chem. 244, 1552–1562 (1969).

    CAS  PubMed  Google Scholar 

  110. Menzies, R. A. & Gold, P. H. The turnover of mitochondria in a variety of tissues of young adult and aged rats. J. Biol. Chem. 246, 2425–2429 (1971).

    CAS  PubMed  Google Scholar 

  111. Chen, H. & Chan, D. C. Mitochondrial dynamics — fusion, fission, movement, and mitophagy — in neurodegenerative diseases. Hum. Mol. Genet. 18, 169–176 (2009).

    Article  CAS  Google Scholar 

  112. Ishihara, N. et al. Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nature Cell Biol. 11, 958–966 (2009).

    Article  CAS  PubMed  Google Scholar 

  113. Liu, X. & Hajnoczky, G. Ca2+-dependent regulation of mitochondrial dynamics by the Miro–Milton complex. Int. J. Biochem . Cell Biol. 41, 1972–1976 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  114. Varadi, A. et al. Cytoplasmic dynein regulates the subcellular distribution of mitochondria by controlling the recruitment of the fission factor dynamin-related protein-1. J. Cell Sci. 117, 4389–4400 (2004).

    Article  CAS  PubMed  Google Scholar 

  115. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  116. Chen, H. et al. Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J. Cell Biol. 160, 189–200 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  117. Detmer, S. A., Vande Velde, C., Cleveland, D. W. & Chan, D. C. Hindlimb gait defects due to motor axon loss and reduced distal muscles in a transgenic mouse model of Charcot-Marie-Tooth type 2A. Hum. Mol. Genet. 17, 367–375 (2008).

    Article  CAS  PubMed  Google Scholar 

  118. Chen, H., McCaffery, J. M. & Chan, D. C. Mitochondrial fusion protects against neurodegeneration in the cerebellum. Cell 130, 548–562 (2007).

    Article  CAS  PubMed  Google Scholar 

  119. Weihofen, A. et al. Pink1 forms a multiprotein complex with Miro and Milton, linking Pink1 function to mitochondrial trafficking. Biochemistry 48, 2045–2052 (2009).

    Article  CAS  PubMed  Google Scholar 

  120. Yu, W., Sun, Y., Guo, S. & Lu, B. The PINK1/Parkin pathway regulates mitochondrial dynamics and function in mammalian hippocampal and dopaminergic neurons. Hum. Mol. Genet. 20, 3227–3240 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  121. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  122. Narendra, D., Kane, L. A., Hauser, D. N., Fearnley, I. M. & Youle, R. J. p62/SQSTM1 is required for Parkin-induced mitochondrial clustering but not mitophagy; VDAC1 is dispensable for both. Autophagy 6, 1090–1106 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  124. Geisler, S. et al. PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nature Cell Biol. 12, 119–131 (2010).

    Article  CAS  PubMed  Google Scholar 

  125. Lee, J. Y., Nagano, Y., Taylor, J. P., Lim, K. L. & Yao, T. P. Disease-causing mutations in Parkin impair mitochondrial ubiquitination, aggregation, and HDAC6-dependent mitophagy. J. Cell Biol. 189, 671–679 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  126. Matsuda, N. et al. PINK1 stabilized by mitochondrial depolarization recruits Parkin to damaged mitochondria and activates latent Parkin for mitophagy. J. Cell Biol. 189, 211–221 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  127. Vives-Bauza, C. et al. PINK1-dependent recruitment of Parkin to mitochondria in mitophagy. Proc. Natl Acad. Sci. USA 107, 378–383 (2010).

    Article  CAS  PubMed  Google Scholar 

  128. Chan, N. C. et al. Broad activation of the ubiquitin-proteasome system by Parkin is critical for mitophagy. Hum. Mol. Genet. 20, 1726–1737 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Yoshii, S. R., Kishi, C., Ishihara, N. & Mizushima, N. Parkin mediates proteasome-dependent protein degradation and rupture of the outer mitochondrial membrane. J. Biol. Chem. 286, 19630–19640 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  130. Wang, X. et al. PINK1 and Parkin target Miro for phosphorylation and degradation to arrest mitochondrial motility. Cell 147, 893–906 (2011). This study demonstrated that the PINK1–parkin pathway also regulates mitochondrial transport, which may help to quarantine parkin-labelled damaged mitochondria for clearance by mitophagy.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  131. Deng, H., Dodson, M. W., Huang, H. & Guo, M. The Parkinson's disease genes pink1 and parkin promote mitochondrial fission and/or inhibit fusion in Drosophila. Proc. Natl Acad. Sci. USA 105, 14503–14508 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  132. Yang, Y. et al. Pink1 regulates mitochondrial dynamics through interaction with the fission/fusion machinery. Proc. Natl Acad. Sci. USA 105, 7070–7075 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Poole, A. C. et al. The PINK1/Parkin pathway regulates mitochondrial morphology. Proc. Natl Acad. Sci. USA 105, 1638–1643 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  134. 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).

    Article  CAS  PubMed  Google Scholar 

  135. Mortiboys, H. et al. Mitochondrial function and morphology are impaired in parkin-mutant fibroblasts. Ann. Neurol. 64, 555–565 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Exner, N. et al. Loss-of-function of human PINK1 results in mitochondrial pathology and can be rescued by parkin. J. Neurosci. 27, 12413–12418 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  137. Wood-Kaczmar, A. et al. PINK1 is necessary for long term survival and mitochondrial function in human dopaminergic neurons. PLoS ONE 3, e2455 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  138. Dagda, R. K. et al. Loss of PINK1 function promotes mitophagy through effects on oxidative stress and mitochondrial fission. J. Biol. Chem. 284, 13843–13855 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  139. Sandebring, A. et al. Mitochondrial alterations in PINK1 deficient cells are influenced by calcineurin-dependent dephosphorylation of dynamin-related protein 1. PLoS ONE 4, e5701 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  140. Lutz, A. K. et al. Loss of parkin or PINK1 function increases Drp1-dependent mitochondrial fragmentation. J. Biol. Chem. 284, 22938–22951 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  141. Ziviani, E., Tao, R. N. & Whitworth, A. J. Drosphila Parkin requires PINK1 for mitochondrial translocation and ubiquitinates Mitofusin. Proc. Natl Acad. Sci. USA 107, 5018–5023 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  142. Gegg, M. E. et al. Mitofusin 1 and mitofusin 2 are ubiquitinated in a PINK1/parkin-dependent manner upon induction of mitophagy. Hum. Mol. Genet. 19, 4861–4870 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  143. Poole, A. C. et al. The mitochondrial fusion-promoting factor mitofusin is a substrate of the PINK1/parkin pathway. PLoS ONE 5, e10054 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  144. Tanaka, A. et al. Proteasome and p97 mediate mitophagy and degradation of mitofusins induced by Parkin. J. Cell Biol. 191, 1367–1380 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  145. Miller, K. E. & Sheetz, M. P. Axonal mitochondrial transport and potential are correlated. J. Cell Sci. 117, 2791–2804 (2004). This study demonstrates that mitochondria with a high membrane potential are transported anterogradely towards distal processes, whereas damaged mitochondria return to the cell body following acute depolarization.

    Article  CAS  PubMed  Google Scholar 

  146. Katsumata, K. et al. Dynein- and activity-dependent retrograde transport of autophagosomes in neuronal axons. Autophagy 6, 378–385 (2010).

    Article  CAS  PubMed  Google Scholar 

  147. Cai, Q. & Sheng, Z. H. Uncovering the role of Snapin in regulating autophagy-lysosomal function. Autophagy 7, 445–447 (2011).

    Article  PubMed  PubMed Central  Google Scholar 

  148. Lee, S., Sato, Y. & Nixon, R. A. Lysosomal proteolysis inhibition selectively disrupts axonal transport of degradative organelles and causes an Alzheimer's-like axonal dystrophy. J. Neurosci. 31, 7817–7830 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  149. Verburg, J. & Hollenbeck, P. J. Mitochondrial membrane potential in axons increases with local nerve growth factor or semaphorin signaling. J. Neurosci. 28, 8306–8315 (2008). This paper provides evidence that mobile and stationary mitochondria show no difference in membrane potential under physiological conditions.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  150. Green, D. R. & Houten, B. V. SnapShot: mitochondrial quality control. Cell 147, 950–950e1 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Karki, S. & Holzbaur, E. L. Cytoplasmic dynein and dynactin in cell division and intracellular transport. Curr. Opin. Cell Biol. 11, 45–53 (1999).

    Article  CAS  PubMed  Google Scholar 

  152. Foth, B. J., Goedecke, M. C. & Soldati, D. New insights into myosin evolution and classification. Proc. Natl Acad. Sci. USA 103, 3681–3686 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  153. Cai, Q. & Sheng, Z.-H. in Mitochondrial Dynamics and Neurodegeneration (ed. Lu, B.) 139–168 (Springer, Dordrecht, 2011).

    Book  Google Scholar 

  154. de Castro, I. P., Martins, L. M. & Tufi, R. Mitochondrial quality control and neurological disease: an emerging connection. Expert Rev. Mol. Med. 19, e12 (2010).

    Article  CAS  Google Scholar 

  155. Tatsuta, T. & Langer, T. Quality control of mitochondria: protection against neurodegeneration and ageing. EMBO J. 27, 306–314 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  156. Detmer, S. A. & Chan, D. C. Functions and dysfunctions of mitochondrial dynamics. Nature Rev. Mol. Cell Biol. 8, 870–879 (2007).

    Article  CAS  Google Scholar 

  157. Westermann, B. Mitochondrial fusion and fission in cell life and death. Nature Rev. Mol. Cell Biol. 11, 872–884 (2010).

    Article  CAS  Google Scholar 

  158. Youle, R. J. & Narendra, D. P. Mechanisms of mitophagy. Nature Rev. Mol. Cell Biol. 12, 9–14 (2011).

    Article  CAS  Google Scholar 

  159. Wang, X. et al. The role of abnormal mitochondrial dynamics in the pathogenesis of Alzheimer's disease. J. Neurochem. 109, 153–159 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  160. Rui, Y., Tiwari, P., Xie, Z. & Zheng, J. Q. Acute impairment of mitochondrial trafficking by β-amyloid peptides in hippocampal neurons. J. Neurosci. 26, 10480–10487 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  161. Stokin, G. B. et al. Axonopathy and transport deficits early in the pathogenesis of Alzheimer's disease. Science 307, 1282–1288 (2005).

    Article  CAS  PubMed  Google Scholar 

  162. Sasaki, S. & Iwata, M. Impairment of fast axonal transport in the proximal axons of anterior horn neurons in amyotrophic lateral sclerosis. Neurology 47, 535–540 (1996).

    Article  CAS  PubMed  Google Scholar 

  163. De Vos, K. J. et al. Familial amyotrophic lateral sclerosis-linked SOD1 mutants perturb fast axonal transport to reduce axonal mitochondria content. Hum. Mol. Genet. 16, 2720–2728 (2007).

    Article  CAS  PubMed  Google Scholar 

  164. Magrané, J. et al. Mutant SOD1 in neuronal mitochondria causes toxicity and mitochondrial dynamics abnormalities. Hum. Mol. Genet. 18, 4552–4564 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  165. Shi, P., Ström, A. L., Gal, J. & Zhu, H. Effects of ALS-related SOD1 mutants on dynein- and KIF5-mediated retrograde and anterograde axonal transport. Biochim. Biophys. Acta 1802, 707–716 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Millecamps, S. et al. Alsin is partially associated with centrosome in human cells. Biochim. Biophys. Acta 1745, 84–100 (2005).

    Article  CAS  PubMed  Google Scholar 

  167. Shan, X., Chiang, P. M., Price, D. L. & Wong, P. C. Altered distributions of Gemini of coiled bodies and mitochondria in motor neurons of TDP-43 transgenic mice. Proc. Natl Acad. Sci. USA 107, 16325–16330 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  168. Bosco, D. A. et al. Wild-type and mutant SOD1 share an aberrant conformation and a common pathogenic pathway in ALS. Nature Neurosci. 13, 1396–1403 (2010).

    Article  CAS  PubMed  Google Scholar 

  169. Zhu, Y. B. & Sheng, Z. H. Increased axonal mitochondrial mobility does not slow amyotrophic lateral sclerosis (ALS)-like disease in mutant SOD1 mice. J. Biol. Chem. 286, 23432–23440 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  170. Caviston, J. P. & Holzbaur, E. L. Huntingtin as an essential integrator of intracellular vesicular trafficking. Trends Cell Biol. 19, 147–155 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  171. Caviston, J. P. et al. Huntingtin facilitates dynein/dynactin-mediated vesicle transport. Proc. Natl Acad. Sci. USA 104, 10045–10050 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Colin, E. et al. Huntingtin phosphorylation acts as a molecular switch for anterograde/retrograde transport in neurons. EMBO J. 27, 2124–2134 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Trushina, E. et al. Mutant huntingtin impairs axonal trafficking in mammalian neurons in vivo and in vitro. Mol. Cell. Biol. 24, 8195–8209 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  174. Chang, D. T., Rintoul, G. L., Pandipati, S. & Reynolds, I. J. Mutant huntingtin aggregates impair mitochondrial movement and trafficking in cortical neurons. Neurobiol. Dis. 22, 388–400 (2006).

    Article  CAS  PubMed  Google Scholar 

  175. Song, W. et al. Mutant huntingtin binds the mitochondrial fission GTPase dynamin-related protein-1 and increases its enzymatic activity. Nature Med. 17, 377–382 (2011).

    Article  CAS  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  177. Lee, H. J., Khoshaghideh, F., Lee, S. & Lee, S. J. Impairment of microtubule-dependent trafficking by overexpression of α-synuclein. Eur . J. Neurosci. 24, 3153–3162 (2006).

    Article  PubMed  Google Scholar 

  178. Yang, F. et al. Parkin stabilizes microtubules through strong binding mediated by three independent domains. J. Biol. Chem. 280, 17154–17162 (2005).

    Article  CAS  PubMed  Google Scholar 

  179. Gillardon, F. Leucine-rich repeat kinase 2 phosphorylates brain tubulin-beta isoforms and modulates microtubule stability — a point of convergence in Parkinsonian neurodegeneration? J. Neurochem. 110, 1514–1522 (2009).

    Article  CAS  PubMed  Google Scholar 

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Acknowledgements

The authors thank M. Davis for editing and other members of the Sheng laboratory for their assistance and discussion. This work was supported by the Intramural Research Program of the US National Institute of Neurological Disorder and Stroke (NINDS), US National Institutes of Health (NIH) (Z.-H.S.) and the NIH Pathway to Independence Award K99 (Q.C.).

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Supplementary Information S1

Representative time-lapse images of axonal mitochondrial motility in wild-type cultured hippocampal neurons. (PDF 90 kb)

Supplementary Information S1 (movie) (MOV 13625 kb)

Supplementary Information S2

Representative time-lapse images of axonal mitochondrial motility in cultured hippocampal neurons from the syntaphilin−/− mice. (PDF 106 kb)

Supplementary Information S2 (movie) (MOV 12310 kb)

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Glossary

Nodes of Ranvier

Regularly spaced gaps in the myelin sheath that surrounds myelinated axons. They expose the axonal plasma membrane to the extracellular fluid. Nodes of Ranvier contain large numbers of voltage-gated ion channels and thus enable conduction of the action potential.

Autophagy–lysosomal system

A primary cellular route for the breakdown of organelles and the degradation of cytoplasmic components. Following the sequestration of organelles and cytoplasm within a double-membrane-bound vacuole (autophagosome), fusion with lysosomes occurs. Lysosomal hydrolases in these 'autolysosomes' degrade their contents.

EF hand

A Ca2+-binding domain that was originally identified in parvalbumin. EF hands are also known as helix–turn–helix domains.

Small interfering RNA

(siRNA). A sequence-specific gene-silencing tool used in RNA interference. siRNAs are short fragments of synthetic double-stranded RNA with 21–23 pairs of nucleotides that have sequence specificity to the gene of interest (the target). These small double-stranded RNAs trigger degradation of the target RNA, thereby creating a partial loss-of-function.

RNA interference

(RNAi). A method by which double-stranded RNA that is encoded in an exogenous vector can be used to interfere with normal RNA processing, causing rapid degradation of the endogenous RNA and thereby precluding translation. This provides a simple way of studying the effects of the absence of a gene product in simple organisms and cells.

Processivity

Motor proteins move stepwise along microtubules without detachment over long distances at the expense of ATP. This movement, termed hand-over-hand motility, is based on the coordinated action of two motor heads that bind one after another to microtubules. This mechanism requires precise coordination of the microtubule affinity of the two motor domains.

Saltatory mobility patterns

The complex nature of mitochondrial transport along neuronal processes, whereby mitochondria move bidirectionally, pause and start moving again, slow down and speed up, and frequently change direction.

Na+/K+ ATPases

Also known as Na+/K+ pumps, these membrane proteins use ATP hydrolysis to move Na+ and K+ in opposite directions across the plasma membrane. They are responsible for maintaining transmembrane concentration gradients for both Na+ and K+ and have a particularly important role in enabling neurons to respond to stimuli and transmit impulses.

Short-term facilitation

A transient increase in synaptic strength occurs when two or more action potentials invade the presynaptic terminal in close succession or as a result of a high-frequency burst of presynaptic action potentials. Facilitation results in more neurotransmitter being released in response to each succeeding action potential owing to prolonged elevation of presynaptic Ca2+ levels following synaptic activity.

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Sheng, ZH., Cai, Q. Mitochondrial transport in neurons: impact on synaptic homeostasis and neurodegeneration. Nat Rev Neurosci 13, 77–93 (2012). https://doi.org/10.1038/nrn3156

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