Axonal Organelles as Molecular Platforms for Axon Growth and Regeneration after Injury
Abstract
:1. Introduction
2. Endoplasmic Reticulum
2.1. Structure and Function
2.2. ER Shaping and Distribution in Axon Growth and Regeneration
2.3. ER in Lipid Synthesis and Trafficking during Axon Growth and Regeneration
2.4. ER in Protein Synthesis and Trafficking during Axon Growth and Regeneration
2.5. ER Calcium Buffering during Axon Growth and Regeneration
3. Mitochondria
3.1. Structure and Function
3.2. Mitochondrial Fission and Fusion in Axon Growth and Regeneration
3.3. Mitochondrial Transport in Axon Growth and Regeneration
3.4. Mitochondrial Calcium Dynamics in Axon Growth and Regeneration
3.5. Mitochondria as Molecular Platforms for Axon Growth and Regeneration
4. Endosomes
4.1. Structure and Function
4.2. Endosomal Regulation by Rab11 in Axon Growth and Regeneration
4.3. Endosome Regulation by Other Rabs in Axon Growth and Regeneration
5. Lysosomes and Autophagosomes
5.1. Structure and Function
5.2. Lysosomal Regulation of Autophagy in Axon Growth and Regeneration
5.3. Lysosome/Endosome Regulation of Intracellular Pathways in Axon Growth and Regeneration
5.4. Lysosome/Late Endosome Regulation of Exocytosis for Axon Growth and Regeneration
6. Proteasome
6.1. Structure and Function
6.2. Protein Degradation by the Proteasome in Developmental Axon Growth
6.3. Protein Degradation by the Proteasome in Axon Regeneration
7. Organellar Interconnections
7.1. ER-Mitochondria Interactions
7.2. ER-Lysosomes/Late Endosomes Interactions
7.3. Endosome/Lysosome-Mitochondria Interactions
7.4. Proteasome-Other Organelles
8. Conclusions
Author Contributions
Funding
Conflicts of Interest
References
- Banker, G. The development of neuronal polarity: A retrospective view. J. Neurosci. 2018, 38, 1867–1873. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Schelski, M.; Bradke, F. Neuronal polarization: From spatiotemporal signaling to cytoskeletal dynamics. Mol. Cell. Neurosci. 2017, 84, 11–28. [Google Scholar] [CrossRef] [PubMed]
- Dotti, C.G.; Sullivan, C.A.; Banker, G.A. The establishment of polarity by hippocampal neurons in culture. J. Neurosci. 1988, 8, 1454–1468. [Google Scholar] [CrossRef] [Green Version]
- Esch, T.; Lemmon, V.; Banker, G. Local presentation of substrate molecules directs axon specification by cultured hippocampal neurons. J. Neurosci. 1999, 19, 6417–6426. [Google Scholar] [CrossRef]
- O’Donnell, M.; Chance, R.K.; Bashaw, G.J. Axon Growth and Guidance: Receptor Regulation and Signal Transduction. Annu. Rev. Neurosci. 2009, 32, 383–412. [Google Scholar] [CrossRef] [Green Version]
- Pfenninger, K.H. Plasma membrane expansion: A neuron’s Herculean task. Nat. Rev. Neurosci. 2009, 10, 251–261. [Google Scholar] [CrossRef] [PubMed]
- Shen, K.; Scheiffele, P. Genetics and cell biology of building specific synaptic connectivity. Annu. Rev. Neurosci. 2010, 33, 473–507. [Google Scholar] [CrossRef] [Green Version]
- Lu, Y.; Brommer, B.; Tian, X.; Krishnan, A.; Meer, M.; Wang, C.; Vera, D.L.; Zeng, Q.; Yu, D.; Bonkowski, M.S.; et al. Reprogramming to recover youthful epigenetic information and restore vision. Nature 2020, 588, 124–129. [Google Scholar] [CrossRef]
- Hilton, B.J.; Bradke, F. Can injured adult CNS axons regenerate by recapitulating development? Development 2017, 144, 3417–3429. [Google Scholar] [CrossRef] [Green Version]
- He, Z.; Jin, Y. Intrinsic Control of Axon Regeneration. Neuron 2016, 90, 437–451. [Google Scholar] [CrossRef] [Green Version]
- Kiyoshi, C.; Tedeschi, A. Axon growth and synaptic function: A balancing act for axonal regeneration and neuronal circuit formation in CNS trauma and disease. Dev. Neurobiol. 2020, 80, 277–301. [Google Scholar] [CrossRef]
- Petrova, V.; Eva, R. The Virtuous Cycle of Axon Growth: Axonal Transport of Growth-Promoting Machinery as an Intrinsic Determinant of Axon Regeneration. Dev. Neurobiol. 2018, 78, 898–925. [Google Scholar] [CrossRef]
- Schwab, M.E.; Strittmatter, S.M. Nogo limits neural plasticity and recovery from injury. Curr. Opin. Neurobiol. 2014, 27, 53–60. [Google Scholar] [CrossRef] [Green Version]
- Bradke, F.; Fawcett, J.W.; Spira, M.E. Assembly of a new growth cone after axotomy: The precursor to axon regeneration. Nat. Rev. Neurosci. 2012, 13, 183–193. [Google Scholar] [CrossRef] [PubMed]
- Ziv, N.E.; Spira, M.E. Localized and transient elevations of intracellular Ca2+ induce the dedifferentiation of axonal segments into growth cones. J. Neurosci. 1997, 17, 3568–3579. [Google Scholar] [CrossRef] [Green Version]
- Chandran, V.; Coppola, G.; Nawabi, H.; Omura, T.; Versano, R.; Huebner, E.A.; Zhang, A.; Costigan, M.; Yekkirala, A.; Barrett, L.; et al. A Systems-Level Analysis of the Peripheral Nerve Intrinsic Axonal Growth Program. Neuron 2016, 89, 956–970. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Costigan, M.; Befort, K.; Karchewski, L.; Griffin, R.S.; D’Urso, D.; Allchorne, A.; Sitarski, J.; Mannion, J.W.; Pratt, R.E.; Woolf, C.J. Replicate high-density rat genome oligonucleotide microarrays reveal hundreds of regulated genes in the dorsal root ganglion after peripheral nerve injury. BMC Neurosci. 2002, 3, 16. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Park, K.K.; Liu, K.; Hu, Y.; Smith, P.D.; Wang, C.; Cai, B.; Xu, B.; Connolly, L.; Kramvis, I.; Sahin, M.; et al. Promoting Axon Regeneration in the Adult CNS by Modulation of the PTEN/mTOR Pathway. Science 2008, 322, 963–966. [Google Scholar] [CrossRef] [Green Version]
- Liu, K.; Lu, Y.; Lee, J.K.; Samara, R.; Willenberg, R.; Sears-Kraxberger, I.; Tedeschi, A.; Park, K.K.; Jin, D.; Cai, B.; et al. PTEN deletion enhances the regenerative ability of adult corticospinal neurons. Nat. Neurosci. 2010, 13, 1075–1081. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Nieuwenhuis, B.; Barber, A.C.; Evans, R.S.; Pearson, C.S.; Fuchs, J.; MacQueen, A.R.; Erp, S.; Haenzi, B.; Hulshof, L.A.; Osborne, A.; et al. PI 3-kinase delta enhances axonal PIP3 to support axon regeneration in the adult CNS. EMBO Mol. Med. 2020, 12, e11674. [Google Scholar] [CrossRef]
- Sun, F.; Park, K.K.; Belin, S.; Wang, D.; Lu, T.; Chen, G.; Zhang, K.; Yeung, C.; Feng, G.; Yankner, B.A.; et al. Sustained axon regeneration induced by co-deletion of PTEN and SOCS3. Nature 2011, 480, 372–375. [Google Scholar] [CrossRef] [Green Version]
- Smith, P.D.; Sun, F.; Park, K.K.; Cai, B.; Wang, C.; Kuwako, K.; Martinez-Carrasco, I.; Connolly, L.; He, Z. SOCS3 Deletion Promotes Optic Nerve Regeneration In Vivo. Neuron 2009, 64, 617–623. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Dupraz, S.; Grassi, D.; Bernis, M.E.; Sosa, L.; Bisbal, M.; Gastaldi, L.; Jausoro, I.; Caceres, A.; Pfenninger, K.H.; Quiroga, S. The TC10-Exo70 Complex Is Essential for Membrane Expansion and Axonal Specification in Developing Neurons. J. Neurosci. 2009, 29, 13292–13301. [Google Scholar] [CrossRef] [Green Version]
- Porter, K.R.; Blum, J. A study in microtomy for electron microscopy. Anat. Rec. 1953, 117, 685–709. [Google Scholar] [CrossRef] [PubMed]
- Nixon-Abell, J.; Obara, C.J.; Weigel, A.V.; Li, D.; Legant, W.R.; Xu, C.S.; Pasolli, H.A.; Harvey, K.; Hess, H.F.; Betzig, E.; et al. Increased spatiotemporal resolution reveals highly dynamic dense tubular matrices in the peripheral ER. Science 2016, 354, aaf3928. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- King, C.; Sengupta, P.; Seo, A.Y.; Lippincott-Schwartz, J. ER membranes exhibit phase behavior at sites of organelle contact. Proc. Natl. Acad. Sci. USA 2020, 117, 7225–7235. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Tsukita, S.; Ishikawa, H. Three-Dimensional Distributionof Smooth Endoplasmic Reticulum in Myelinated Axons. J. Electron Microsc. 1976, 25, 141–149. [Google Scholar]
- Droz, B.; Rambourg, A.; Koenig, H.L. The smooth endoplasmic reticulum: Structure and role in the renewal of axonal membrane and synaptic vesicles by fast axonal tranport. Brain Res. 1975, 93, 1–13. [Google Scholar] [CrossRef]
- González, C.; Cánovas, J.; Fresno, J.; Couve, E.; Court, F.A.; Couve, A. Axons provide the secretory machinery for trafficking of voltage-gated sodium channels in peripheral nerve. Proc. Natl. Acad. Sci. USA 2016, 113, 1823–1828. [Google Scholar] [CrossRef] [Green Version]
- Cioni, J.-M.; Lin, J.Q.; Holtermann, A.V.; Koppers, M.; Jakobs, M.A.H.; Azizi, A.; Turner-Bridger, B.; Shigeoka, T.; Franze, K.; Harris, W.A.; et al. Late Endosomes Act as mRNA Translation Platforms and Sustain Mitochondria in Axons. Cell 2019, 176, 56–72.e15. [Google Scholar] [CrossRef] [Green Version]
- Wu, Y.; Whiteus, C.; Xu, C.S.; Hayworth, K.J.; Weinberg, R.J.; Hess, H.F.; De Camilli, P. Contacts between the endoplasmic reticulum and other membranes in neurons. Proc. Natl. Acad. Sci. USA 2017, 114, E4859–E4867. [Google Scholar] [CrossRef] [Green Version]
- Öztürk, Z.; O’Kane, C.J.; Pérez-Moreno, J.J. Axonal Endoplasmic Reticulum Dynamics and Its Roles in Neurodegeneration. Front. Neurosci. 2020, 14, 48. [Google Scholar] [CrossRef] [PubMed]
- Westrate, L.M.; Lee, J.E.; Prinz, W.A.; Voeltz, G.K. Form follows function: The importance of endoplasmic reticulum shape. Annu. Rev. Biochem. 2015, 84, 791–811. [Google Scholar] [CrossRef]
- Zhang, H.; Hu, J. Shaping the Endoplasmic Reticulum into a Social Network. Trends Cell Biol. 2016, 26, 934–943. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Hübner, C.A.; Kurth, I. Membrane-shaping disorders: A common pathway in axon degeneration. Brain 2014, 137, 3109–3121. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Blackstone, C. Cellular Pathways of Hereditary Spastic Paraplegia. Annu. Rev. Neurosci. 2012, 35, 25–47. [Google Scholar] [CrossRef] [Green Version]
- Blackstone, C.; O’Kane, C.J.; Reid, E. Hereditary spastic paraplegias: Membrane traffic and the motor pathway. Nat. Rev. Neurosci. 2011, 12, 31–42. [Google Scholar] [CrossRef] [Green Version]
- Yalçın, B.; Zhao, L.; Stofanko, M.; O’Sullivan, N.C.; Kang, Z.H.; Roost, A.; Thomas, M.R.; Zaessinger, S.; Blard, O.; Patto, A.L.; et al. Modeling of axonal endoplasmic reticulum network by spastic paraplegia proteins. Elife 2017, 6, e23882. [Google Scholar] [CrossRef]
- O’Sullivan, N.C.; Jahn, T.R.; Reid, E.; O’Kane, C.J. Reticulon-like-1, the Drosophila orthologue of the Hereditary Spastic Paraplegia gene reticulon 2, is required for organization of endoplasmic reticulum and of distal motor axons. Hum. Mol. Genet. 2012, 21, 3356–3365. [Google Scholar] [CrossRef] [Green Version]
- Oliva, M.K.; Pérez-Moreno, J.J.; O’Shaughnessy, J.; Wardill, T.J.; O’Kane, C.J. Endoplasmic Reticulum Lumenal Indicators in Drosophila Reveal Effects of HSP-Related Mutations on Endoplasmic Reticulum Calcium Dynamics. Front. Neurosci. 2020, 14, 816. [Google Scholar] [CrossRef]
- English, A.R.; Voeltz, G.K. Endoplasmic reticulum structure and interconnections with other organelles. Cold Spring Harb. Perspect. Biol. 2013, 5, 1–16. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Farías, G.G.; Fréal, A.; Tortosa, E.; Stucchi, R.; Pan, X.; Portegies, S.; Will, L.; Altelaar, M.; Hoogenraad, C.C. Feedback-Driven Mechanisms between Microtubules and the Endoplasmic Reticulum Instruct Neuronal Polarity. Neuron 2019, 102, 184–201.e8. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Petrova, V.; Pearson, C.S.; Ching, J.; Tribble, J.R.; Solano, A.G.; Yang, Y.; Love, F.M.; Watt, R.J.; Osborne, A.; Reid, E.; et al. Protrudin functions from the endoplasmic reticulum to support axon regeneration in the adult CNS. Nat. Commun. 2020, 11, 5614. [Google Scholar] [CrossRef] [PubMed]
- Park, S.H.; Zhu, P.P.; Parker, R.L.; Blackstone, C. Hereditary spastic paraplegia proteins REEP1, spastin, and atlastin-1 coordinate microtubule interactions with the tubular ER network. J. Clin. Investig. 2010, 120, 1097–1110. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Lim, Y.; Cho, I.T.; Schoel, L.J.; Cho, G.; Golden, J.A. Hereditary spastic paraplegia-linked REEP1 modulates endoplasmic reticulum/mitochondria contacts. Ann. Neurol. 2015, 78, 679–696. [Google Scholar] [CrossRef] [PubMed]
- Rao, K.; Stone, M.C.; Weiner, A.T.; Gheres, K.W.; Zhou, C.; Deitcher, D.L.; Levitan, E.S.; Rolls, M.M. Spastin, atlastin, and ER relocalization are involved in axon but not dendrite regeneration. Mol. Biol. Cell 2016, 27, 3245–3256. [Google Scholar] [CrossRef]
- Shorey, M.; Stone, M.C.; Mandel, J.; Rolls, M.M. Neurons survive simultaneous injury to axons and dendrites and regrow both types of processes in vivo. Dev. Biol. 2020, 465, 108–118. [Google Scholar] [CrossRef]
- Zhu, P.P.; Soderblom, C.; Tao-Cheng, J.H.; Stadler, J.; Blackstone, C. SPG3A protein atlastin-1 is enriched in growth cones and promotes axon elongation during neuronal development. Hum. Mol. Genet. 2006, 15, 1343–1353. [Google Scholar] [CrossRef] [Green Version]
- Nozumi, M.; Togano, T.; Takahashi-Niki, K.; Lu, J.; Honda, A.; Taoka, M.; Shinkawa, T.; Koga, H.; Takeuchi, K.; Isobe, T.; et al. Identification of functional marker proteins in the mammalian growth cone. Proc. Natl. Acad. Sci. USA 2009, 106, 17211–17216. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Saita, S.; Shirane, M.; Natume, T.; Iemura, S.; Nakayama, K.I. Promotion of Neurite Extension by Protrudin Requires Its Interaction with Vesicle-associated Membrane Protein-associated Protein. J. Biol. Chem. 2009, 284, 13766–13777. [Google Scholar] [CrossRef] [Green Version]
- De Vincentiis, S.; Falconieri, A.; Mainardi, M.; Cappello, V.; Scribano, V.; Bizzarri, R.; Storti, B.; Dente, L.; Costa, M.; Raffa, V. Extremely Low Forces Induce Extreme Axon Growth. J. Neurosci. 2020, 40, 4997–5007. [Google Scholar] [CrossRef] [PubMed]
- Vance, J.E. Newly made phosphatidylserine and phosphatidylethanolamine are preferentially translocated between rat liver mitochondria and endoplasmic reticulum. J. Biol. Chem. 1991, 266, 89–97. [Google Scholar] [CrossRef]
- Vance, J.E.; Pan, D.; Campenot, R.B.; Bussiere, M.; Vance, D.E. Evidence that the Major Membrane Lipids, Except Cholesterol, Are Made in Axons of Cultured Rat Sympathetic Neurons. J. Neurochem. 2008, 62, 329–337. [Google Scholar] [CrossRef] [PubMed]
- De Chaves, E.P.; Vance, D.E.; Campenot, R.B.; Vance, J.E. Axonal synthesis of phosphatidylcholine is required for normal axonal growth in rat sympathetic neurons. J. Cell Biol. 1995, 128, 913–918. [Google Scholar] [CrossRef] [PubMed]
- Luarte, A.; Cornejo, V.H.; Bertin, F.; Gallardo, J.; Couve, A. The axonal endoplasmic reticulum: One organelle—many functions in development, maintenance, and plasticity. Dev. Neurobiol. 2018, 78, 181–208. [Google Scholar] [CrossRef] [PubMed]
- Jacquemyn, J.; Cascalho, A.; Goodchild, R.E. The ins and outs of endoplasmic reticulum-controlled lipid biosynthesis. EMBO Rep. 2017, 18, 1905–1921. [Google Scholar] [CrossRef] [PubMed]
- Rodríguez-Berdini, L.; Orlando Ferrero, G.; Bustos Plonka, F.; Mauricio Cardozo Gizzi, A.; Prucca, C.G.; Quiroga, S.; Leonor Caputto, B.; Orio, O.A. The moonlighting protein c-Fos activates lipid synthesis in neurons, an activity that is critical for cellular differentiation and cortical development. J. Biol. Chem. 2020, 295, 8808–8818. [Google Scholar] [CrossRef]
- Bakan, I.; Laplante, M. Connecting mTORC1 signaling to SREBP-1 activation. Curr. Opin. Lipidol. 2012, 23, 226–234. [Google Scholar] [CrossRef]
- Ricoult, S.J.H.; Yecies, J.L.; Ben-Sahra, I.; Manning, B.D. Oncogenic PI3K and K-Ras stimulate de novo lipid synthesis through mTORC1 and SREBP. Oncogene 2016, 35, 1250–1260. [Google Scholar] [CrossRef] [Green Version]
- Ricoult, S.J.H.; Manning, B.D. The multifaceted role of mTORC1 in the control of lipid metabolism. EMBO Rep. 2013, 14, 242–251. [Google Scholar] [CrossRef] [Green Version]
- Lev, S. Nonvesicular lipid transfer from the endoplasmic reticulum. Cold Spring Harb. Perspect. Biol. 2012, 4, a013300. [Google Scholar] [CrossRef]
- Lev, S. Non-vesicular lipid transport by lipid-transfer proteins and beyond. Nat. Rev. Mol. Cell Biol. 2010, 11, 739–750. [Google Scholar] [CrossRef]
- Wojnacki, J.; Galli, T. Membrane traffic during axon development. Dev. Neurobiol. 2016, 76, 1185–1200. [Google Scholar] [CrossRef] [PubMed]
- Kaplan, M.R.; Simoni, R.D. Intracellular transport of phosphatidylcholine to the plasma membrane. J. Cell Biol. 1985, 101, 441–445. [Google Scholar] [CrossRef] [PubMed]
- Petkovic, M.; Jemaiel, A.; Daste, F.; Specht, C.G.; Izeddin, I.; Vorkel, D.; Verbavatz, J.-M.; Darzacq, X.; Triller, A.; Pfenninger, K.H.; et al. The SNARE Sec22b has a non-fusogenic function in plasma membrane expansion. Nat. Cell Biol. 2014, 16, 434–444. [Google Scholar] [CrossRef] [PubMed]
- Gallo, A.; Vannier, C.; Galli, T. Endoplasmic Reticulum–Plasma Membrane Associations:Structures and Functions. Annu. Rev. Cell Dev. Biol. 2016, 32, 279–301. [Google Scholar] [CrossRef] [PubMed]
- Gallo, A.; Danglot, L.; Giordano, F.; Hewlett, B.; Binz, T.; Vannier, C.; Galli, T. Role of the Sec22b-E-Syt complex in neurite growth and ramification. J. Cell Sci. 2020, 133, jcs247148. [Google Scholar] [CrossRef] [PubMed]
- Blackmore, M.; Letourneau, P.C. Protein synthesis in distal axons is not required for axon growth in the embryonic spinal cord. Dev. Neurobiol. 2007, 67, 976–986. [Google Scholar] [CrossRef] [PubMed]
- Twiss, J.L.; Fainzilber, M. Ribosomes in axons-scrounging from the neighbors? Trends Cell Biol. 2009, 19, 236–243. [Google Scholar] [CrossRef]
- Tuck, E.; Cavalli, V. Roles of membrane trafficking in nerve repair and regeneration. Commun. Integr. Biol. 2010, 3, 209–214. [Google Scholar] [CrossRef]
- Gumy, L.F.; Tan, C.L.; Fawcett, J.W. The role of local protein synthesis and degradation in axon regeneration. Exp. Neurol. 2010, 223, 28–37. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Willis, D.; Ka, W.L.; Zheng, J.Q.; Chang, J.H.; Smit, A.; Kelly, T.; Merianda, T.T.; Sylvester, J.; Van Minnen, J.; Twiss, J.L. Differential transport and local translation of cytoskeletal, injury-response, and neurodegeneration protein mRNAs in axons. J. Neurosci. 2005, 25, 778–791. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kalinski, A.L.; Sachdeva, R.; Gomes, C.; Lee, S.J.; Shah, Z.; Houle, J.D.; Twiss, J.L. mRNAs and protein synthetic machinery localize into regenerating spinal cord axons when they are provided a substrate that supports growth. J. Neurosci. 2015, 35, 10357–10370. [Google Scholar] [CrossRef] [PubMed]
- Verma, P.; Chierzi, S.; Codd, A.M.; Campbell, D.S.; Meyer, R.L.; Holt, C.E.; Fawcett, J.W. Axonal Protein Synthesis and Degradation Are Necessary for Efficient Growth Cone Regeneration. J. Neurosci. 2005, 25, 331–342. [Google Scholar] [CrossRef] [PubMed]
- Merianda, T.T.; Lin, A.C.; Lam, J.S.Y.; Vuppalanchi, D.; Willis, D.E.; Karin, N.; Holt, C.E.; Twiss, J.L. A functional equivalent of endoplasmic reticulum and Golgi in axons for secretion of locally synthesized proteins. Mol. Cell. Neurosci. 2009, 40, 128–142. [Google Scholar] [CrossRef] [Green Version]
- Van Erp, S.; van Berkel, A.A.; Feenstra, E.M.; Sahoo, P.K.; Wagstaff, L.; Twiss, J.L.; Fawcett, J.W.; Eva, R.; Ffrench-Constant, C. Age-related loss of axonal regeneration is reflected by the level of local translation. Exp. Neurol. 2021, 113594. [Google Scholar] [CrossRef]
- Holt, C.E.; Martin, K.C.; Schuman, E.M. Local translation in neurons: Visualization and function. Nat. Struct. Mol. Biol. 2019, 26, 557–566. [Google Scholar] [CrossRef]
- Vuppalanchi, D.; Merianda, T.T.; Donnelly, C.; Pacheco, A.; Williams, G.; Yoo, S.; Ratan, R.R.; Willis, D.E.; Twiss, J.L. Lysophosphatidic acid differentially regulates axonal mRNA translation through 5’UTR elements. Mol. Cell. Neurosci. 2012, 50, 136–146. [Google Scholar] [CrossRef] [Green Version]
- Ying, Z.; Misra, V.; Verge, V.M.K. Sensing nerve injury at the axonal ER: Activated Luman/CREB3 serves as a novel axonally synthesized retrograde regeneration signal. Proc. Natl. Acad. Sci. USA 2014, 111, 16142–16147. [Google Scholar] [CrossRef] [Green Version]
- Friedman, J.R.; Voeltz, G.K. The ER in 3D: A multifunctional dynamic membrane network. Trends Cell Biol. 2011, 21, 709–717. [Google Scholar] [CrossRef] [Green Version]
- De Gregorio, C.; Delgado, R.; Ibacache, A.; Sierralta, J.; Couve, A. Drosophila Atlastin in motor neurons is required for locomotion and presynaptic function. J. Cell Sci. 2017, 130, 3507–3516. [Google Scholar] [CrossRef] [Green Version]
- Liu, Y.; Vidensky, S.; Ruggiero, A.M.; Maier, S.; Sitte, H.H.; Rothstein, J.D. Reticulon RTN2B regulates trafficking and function of neuronal glutamate transporter EAAC1. J. Biol. Chem. 2008, 283, 6561–6571. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kamemura, K.; Chihara, T. Multiple functions of the ER-resident VAP and its extracellular role in neural development and disease. J. Biochem. 2019, 165, 391–400. [Google Scholar] [CrossRef]
- Esk, C.; Lindenhofer, D.; Haendeler, S.; Wester, R.A.; Pflug, F.; Schroeder, B.; Bagley, J.A.; Elling, U.; Zuber, J.; von Haeseler, A.; et al. A human tissue screen identifies a regulator of ER secretion as a brain size determinant. Science 2020, 370, eabb5390. [Google Scholar] [CrossRef]
- Cui-Wang, T.; Hanus, C.; Cui, T.; Helton, T.; Bourne, J.; Watson, D.; Harris, K.M.; Ehlers, M.D. Local zones of endoplasmic reticulum complexity confine cargo in neuronal dendrites. Cell 2012, 148, 309–321. [Google Scholar] [CrossRef] [Green Version]
- Valenzuela, J.I.; Jaureguiberry-Bravo, M.; Salas, D.A.; Ramírez, O.A.; Cornejo, V.H.; Lu, H.E.; Blanpied, T.A.; Couve, A. Transport along the dendritic endoplasmic reticulum mediates the trafficking of GABAB receptors. J. Cell Sci. 2014, 127, 3382–3395. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Bowen, A.B.; Bourke, A.M.; Hiester, B.G.; Hanus, C.; Kennedy, M.J. Golgi-Independent secretory trafficking through recycling endosomes in neuronal dendrites and spines. Elife 2017, 6, e27362. [Google Scholar] [CrossRef]
- Tojima, T.; Akiyama, H.; Itofusa, R.; Li, Y.; Katayama, H.; Miyawaki, A.; Kamiguchi, H. Attractive axon guidance involves asymmetric membrane transport and exocytosis in the growth cone. Nat. Neurosci. 2007, 10, 58–66. [Google Scholar] [CrossRef] [PubMed]
- Tojima, T.; Hines, J.H.; Henley, J.R.; Kamiguchi, H. Second messengers and membrane trafficking direct and organize growth cone steering. Nat. Rev. Neurosci. 2011, 12, 191–203. [Google Scholar] [CrossRef] [Green Version]
- Wada, F.; Nakata, A.; Tatsu, Y.; Ooashi, N.; Fukuda, T.; Nabetani, T.; Kamiguchi, H. Myosin Va and Endoplasmic Reticulum Calcium Channel Complex Regulates Membrane Export during Axon Guidance. CellReports 2016, 15, 1329–1344. [Google Scholar] [CrossRef] [Green Version]
- Pavez, M.; Thompson, A.C.; Arnott, H.J.; Mitchell, C.B.; D’Atri, I.; Don, E.K.; Chilton, J.K.; Scott, E.K.; Lin, J.Y.; Young, K.M.; et al. STIM1 is required for remodeling of the endoplasmic reticulum and microtubule cytoskeleton in steering growth cones. J. Neurosci. 2019, 39, 5095–5114. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kang, F.; Zhou, M.; Huang, X.; Fan, J.; Wei, L.; Boulanger, J.; Liu, Z.; Salamero, J.; Liu, Y.; Chen, L. E-syt1 Re-arranges STIM1 Clusters to Stabilize Ring-shaped ER-PM Contact Sites and Accelerate Ca 2+ Store Replenishment. Sci. Rep. 2019, 9, 1–11. [Google Scholar]
- Zaman, M.F.; Nenadic, A.; Radojičić, A.; Rosado, A.; Beh, C.T. Sticking With It: ER-PM Membrane Contact Sites as a Coordinating Nexus for Regulating Lipids and Proteins at the Cell Cortex. Front. Cell Dev. Biol. 2020, 8, 675. [Google Scholar] [CrossRef]
- Cho, Y.; Sloutsky, R.; Naegle, K.M.; Cavalli, V. XInjury-Induced HDAC5 nuclear export is essential for axon regeneration. Cell 2013, 155, 894. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Ohtake, Y.; Matsuhisa, K.; Kaneko, M.; Kanemoto, S.; Asada, R.; Imaizumi, K.; Saito, A. Axonal Activation of the Unfolded Protein Response Promotes Axonal Regeneration Following Peripheral Nerve Injury. Neuroscience 2018, 375, 34–48. [Google Scholar] [CrossRef] [PubMed]
- Oñate, M.; Catenaccio, A.; Martínez, G.; Armentano, D.; Parsons, G.; Kerr, B.; Hetz, C.; Court, F.A. Activation of the unfolded protein response promotes axonal regeneration after peripheral nerve injury. Sci. Rep. 2016, 6, 1–14. [Google Scholar] [CrossRef] [Green Version]
- Valenzuela, V.; Collyer, E.; Armentano, D.; Parsons, G.B.; Court, F.A.; Hetz, C. Activation of the unfolded protein response enhances motor recovery after spinal cord injury. Cell Death Dis. 2012, 3, 272. [Google Scholar] [CrossRef]
- Hu, Y.; Park, K.K.; Yang, L.; Wei, X.; Yang, Q.; Cho, K.S.; Thielen, P.; Lee, A.H.; Cartoni, R.; Glimcher, L.H.; et al. Differential effects of unfolded protein response pathways on axon injury-induced death of retinal ganglion cells. Neuron 2012, 73, 445–452. [Google Scholar] [CrossRef] [Green Version]
- Halliday, M.; Mallucci, G.R. Review: Modulating the unfolded protein response to prevent neurodegeneration and enhance memory Modulating the unfolded protein response to prevent neurodegeneration and enhance memory. Neuropathol. Appl. Neurobiol. 2015, 41, 414–427. [Google Scholar] [CrossRef] [Green Version]
- Hetz, C.; Mollereau, B. Disturbance of endoplasmic reticulum proteostasis in neurodegenerative diseases. Nat. Rev. Neurosci. 2014, 15, 233–249. [Google Scholar] [CrossRef]
- De Juan-Sanz, J.; Holt, G.T.; Schreiter, E.R.; de Juan, F.; Kim, D.S.; Ryan, T.A. Axonal Endoplasmic Reticulum Ca2+ Content Controls Release Probability in CNS Nerve Terminals. Neuron 2017, 93, 867–881.e6. [Google Scholar] [CrossRef] [Green Version]
- Lindhout, F.W.; Cao, Y.; Kevenaar, J.T.; Bodzęta, A.; Stucchi, R.; Boumpoutsari, M.M.; Katrukha, E.A.; Altelaar, M.; MacGillavry, H.D.; Hoogenraad, C.C. VAP-SCRN1 interaction regulates dynamic endoplasmic reticulum remodeling and presynaptic function. EMBO J. 2019, 38, e101345. [Google Scholar] [CrossRef]
- Kuijpers, M.; Kochlamazashvili, G.; Stumpf, A.; Puchkov, D.; Swaminathan, A.; Lucht, M.T.; Krause, E.; Maritzen, T.; Schmitz, D.; Haucke, V. Neuronal Autophagy Regulates Presynaptic Neurotransmission by Controlling the Axonal Endoplasmic Reticulum. Neuron 2020, 109, 299–313.e9. [Google Scholar] [CrossRef] [PubMed]
- Ferri, K.F.; Kroemer, G. Organelle-specific initiation of cell death pathways. Nat. Cell Biol. 2001, 3, E255–E263. [Google Scholar] [CrossRef]
- Rizzuto, R.; Pozzan, T. Microdomains of intracellular Ca2+: Molecular determinants and functional consequences. Physiol. Rev. 2006, 86, 369–408. [Google Scholar] [CrossRef] [PubMed]
- Bock, F.J.; Tait, S.W.G. Mitochondria as multifaceted regulators of cell death. Nat. Rev. Mol. Cell Biol. 2020, 21, 85–100. [Google Scholar] [CrossRef]
- 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. 2016, 26, 2602–2608. [Google Scholar] [CrossRef] [Green Version]
- Smit-Rigter, L.; Rajendran, R.; Silva, C.A.; Spierenburg, L.; Groeneweg, F.; Ruimschotel, E.M.; van Versendaal, D.; van der Togt, C.; Eysel, U.T.; Alexander Heimel, J.; et al. Mitochondrial Dynamics in Visual Cortex Are Limited In Vivo and Not Affected by Axonal Structural Plasticity. Curr. Biol. 2016, 26, 2609–2616. [Google Scholar] [CrossRef] [Green Version]
- Mar, F.M.; Simões, A.R.; Leite, S.; Morgado, M.M.; Santos, T.E.; Rodrigo, I.S.; Teixeira, C.A.; Misgeld, T.; Sousa, M.M. CNS axons globally increase axonal transport after peripheral conditioning. J. Neurosci. 2014, 34, 5965–5970. [Google Scholar] [CrossRef] [Green Version]
- Misgeld, T.; Kerschensteiner, M.; Bareyre, F.M.; Burgess, R.W.; Lichtman, J.W. Imaging axonal transport of mitochondria in vivo. Nat. Methods 2007, 4, 559–561. [Google Scholar] [CrossRef]
- O’Donnell, K.C.; Vargas, M.E.; Sagasti, A. Wlds and PGC-1α regulate mitochondrial transport and oxidation state after axonal injury. J. Neurosci. 2013, 33, 14778–14790. [Google Scholar] [CrossRef] [Green Version]
- Han, S.M.; Baig, H.S.; Hammarlund, M. Mitochondria Localize to Injured Axons to Support Regeneration. Neuron 2016, 92, 1308–1323. [Google Scholar] [CrossRef] [Green Version]
- Hirokawa, N.; Noda, Y.; Tanaka, Y.; Niwa, S. Kinesin superfamily motor proteins and intracellular transport. Nat. Rev. Mol. Cell Biol. 2009, 10, 682–696. [Google Scholar] [CrossRef] [PubMed]
- Saxton, W.M.; Hollenbeck, P.J. The axonal transport of mitochondria. J. Cell Sci. 2012, 125, 2095–2104. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Lin, M.Y.; Sheng, Z.H. Regulation of mitochondrial transport in neurons. Exp. Cell Res. 2015, 334, 35–44. [Google Scholar] [CrossRef] [Green Version]
- Schwarz, T.L. Mitochondrial trafficking in neurons. Cold Spring Harb. Perspect. Biol. 2013, 5, a011304. [Google Scholar] [CrossRef] [Green Version]
- Chang, D.T.W.; Reynolds, I.J. Differences in mitochondrial movement and morphology in young and mature primary cortical neurons in culture. Neuroscience 2006, 141, 727–736. [Google Scholar] [CrossRef] [PubMed]
- Zhou, B.; Yu, P.; Lin, M.-Y.; Sun, T.; Chen, Y.; Sheng, Z.-H. Facilitation of axon regeneration by enhancing mitochondrial transport and rescuing energy deficits. J. Cell Biol. 2016, 214, 103–119. [Google Scholar] [CrossRef] [Green Version]
- Kang, J.-S.; Tian, J.-H.; Pan, P.-Y.; Zald, P.; Li, C.; Deng, C.; Sheng, Z.-H. Docking of Axonal Mitochondria by Syntaphilin Controls Their Mobility and Affects Short-Term Facilitation. Cell 2008, 132, 137–148. [Google Scholar] [CrossRef] [Green Version]
- Sun, T.; Qiao, H.; Pan, P.Y.; Chen, Y.; Sheng, Z.H. Motile axonal mitochondria contribute to the variability of presynaptic strength. Cell Rep. 2013, 4, 413–419. [Google Scholar] [CrossRef] [Green Version]
- Steketee, M.B.; Moysidis, S.N.; Weinstein, J.E.; Kreymerman, A.; Silva, J.P.; Iqbal, S.; Goldberg, J.L. Mitochondrial dynamics regulate growth cone motility, guidance, and neurite growth rate in perinatal retinal ganglion cells in vitro. Investig. Ophthalmol. Vis. Sci. 2012, 53, 7402–7411. [Google Scholar] [CrossRef] [PubMed]
- Kreymerman, A.; Buickians, D.N.; Nahmou, M.M.; Tran, T.; Galvao, J.; Wang, Y.; Sun, N.; Bazik, L.; Huynh, S.K.; Cho, I.J.; et al. MTP18 is a Novel Regulator of Mitochondrial Fission in CNS Neuron Development, Axonal Growth, and Injury Responses. Sci. Rep. 2019, 9, 1–13. [Google Scholar] [CrossRef]
- Chen, H.; Detmer, S.A.; Ewald, A.J.; Griffin, E.E.; Fraser, S.E.; Chan, D.C. Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J. Cell Biol. 2003, 160, 189–200. [Google Scholar] [CrossRef]
- Davies, V.J.; Hollins, A.J.; Piechota, M.J.; Yip, W.; Davies, J.R.; White, K.E.; Nicols, P.P.; Boulton, M.E.; Votruba, M. Opa1 deficiency in a mouse model of autosomal dominant optic atrophy impairs mitochondrial morphology, optic nerve structure and visual function. Hum. Mol. Genet. 2007, 16, 1307–1318. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- 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. 2018, 9, 1–15. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kiryu-Seo, S.; Tamada, H.; Kato, Y.; Yasuda, K.; Ishihara, N.; Nomura, M.; Mihara, K.; Kiyama, H. Mitochondrial fission is an acute and adaptive response in injured motor neurons. Sci. Rep. 2016, 6, 1–14. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Mattson, M.P.; Partin, J. Evidence for mitochondrial control of neuronal polarity. J. Neurosci. Res. 1999, 56, 8–20. [Google Scholar] [CrossRef]
- Vaarmann, A.; Mandel, M.; Zeb, A.; Wareski, P.; Liiv, J.; Kuum, M.; Antsov, E.; Liiv, M.; Cagalinec, M.; Choubey, V.; et al. Mitochondrial biogenesis is required for axonal growth. Development 2016, 143, 1981–1992. [Google Scholar] [CrossRef] [Green Version]
- Van Spronsen, M.; Mikhaylova, M.; Lipka, J.; Schlager, M.A.; van den Heuvel, D.J.; Kuijpers, M.; Wulf, P.S.; Keijzer, N.; Demmers, J.; Kapitein, L.C.; et al. TRAK/Milton Motor-Adaptor Proteins Steer Mitochondrial Trafficking to Axons and Dendrites. Neuron 2013, 77, 485–502. [Google Scholar] [CrossRef] [Green Version]
- Kalinski, A.L.; Kar, A.N.; Craver, J.; Tosolini, A.P.; Sleigh, J.N.; Lee, S.J.; Hawthorne, A.; Brito-Vargas, P.; Miller-Randolph, S.; Passino, R.; et al. Deacetylation of Miro1 by HDAC6 blocks mitochondrial transport and mediates axon growth inhibition. J. Cell Biol. 2019, 218, 1871–1890. [Google Scholar] [CrossRef] [Green Version]
- Cavallucci, V.; Bisicchia, E.; Cencioni, M.T.; Ferri, A.; Latini, L.; Nobili, A.; Biamonte, F.; Nazio, F.; Fanelli, F.; Moreno, S.; et al. Acute focal brain damage alters mitochondrial dynamics and autophagy in axotomized neurons. Cell Death Dis. 2014, 5, e1545. [Google Scholar] [CrossRef] [Green Version]
- Sheng, Z.H. The Interplay of Axonal Energy Homeostasis and Mitochondrial Trafficking and Anchoring. Trends Cell Biol. 2017, 27, 403–416. [Google Scholar] [CrossRef] [PubMed]
- Cartoni, R.; Norsworthy, M.W.; Bei, F.; Wang, C.; Li, S.; Zhang, Y.; Gabel, C.V.; Schwarz, T.L.; He, Z. The Mammalian-Specific Protein Armcx1 Regulates Mitochondrial Transport during Axon Regeneration. Neuron 2016, 92, 1294–1307. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Han, Q.; Xie, Y.; Ordaz, J.D.; Huh, A.J.; Huang, N.; Wu, W.; Liu, N.; Chamberlain, K.A.; Sheng, Z.H.; Xu, X.M. Restoring Cellular Energetics Promotes Axonal Regeneration and Functional Recovery after Spinal Cord Injury. Cell Metab. 2020, 31, 623–641.e8. [Google Scholar] [CrossRef] [PubMed]
- Knowlton, W.M.; Hubert, T.; Wu, Z.; Chisholm, A.D.; Jin, Y. A select subset of electron transport chain genes associated with optic atrophy link mitochondria to axon regeneration in Caenorhabditis elegans. Front. Neurosci. 2017, 11, 263. [Google Scholar] [CrossRef] [PubMed]
- Rawson, R.L.; Yam, L.; Weimer, R.M.; Bend, E.G.; Hartwieg, E.; Horvitz, H.R.; Clark, S.G.; Jorgensen, E.M. Axons degenerate in the absence of mitochondria in C. elegans. Curr. Biol. 2014, 24, 760–765. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Tang, N.H.; Kim, K.W.; Xu, S.; Blazie, S.M.; Yee, B.A.; Yeo, G.W.; Jin, Y.; Chisholm, A.D. The mRNA Decay Factor CAR-1/LSM14 Regulates Axon Regeneration via Mitochondrial Calcium Dynamics. Curr. Biol. 2020, 30, 865–876.e7. [Google Scholar] [CrossRef] [Green Version]
- Chen, Y.; Sheng, Z.H. Kinesin-1-syntaphilin coupling mediates activity-dependent regulation of axonal mitochondrial transport. J. Cell Biol. 2013, 202, 351–364. [Google Scholar] [CrossRef]
- Wang, X.; Schwarz, T.L. The Mechanism of Ca2+-Dependent Regulation of Kinesin-Mediated Mitochondrial Motility. Cell 2009, 136, 163–174. [Google Scholar] [CrossRef] [Green Version]
- Courchet, J.; Lewis, T.L.; Lee, S.; Courchet, V.; Liou, D.Y.; Aizawa, S.; Polleux, F. Terminal axon branching is regulated by the LKB1-NUAK1 kinase pathway via presynaptic mitochondrial capture. Cell 2013, 153, 1510. [Google Scholar] [CrossRef] [Green Version]
- Tao, K.; Matsuki, N.; Koyama, R. AMP-activated protein kinase mediates activity-dependent axon branching by recruiting mitochondria to axon. Dev. Neurobiol. 2014, 74, 557–573. [Google Scholar] [CrossRef] [PubMed]
- Mehta, S.T.; Luo, X.; Park, K.K.; Bixby, J.L.; Lemmon, V.P. Hyperactivated Stat3 boosts axon regeneration in the CNS. Exp. Neurol. 2016, 280, 115–120. [Google Scholar] [CrossRef] [Green Version]
- Luo, X.; Ribeiro, M.; Bray, E.R.; Lee, D.H.; Yungher, B.J.; Mehta, S.T.; Thakor, K.A.; Diaz, F.; Lee, J.K.; Moraes, C.T.; et al. Enhanced Transcriptional Activity and Mitochondrial Localization of STAT3 Co-induce Axon Regrowth in the Adult Central Nervous System. Cell Rep. 2016, 15, 398–410. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Fang, Y.; Soares, L.; Teng, X.; Geary, M.; Bonini, N.M. A novel Drosophila model of nerve injury reveals an essential role of Nmnat in maintaining axonal integrity. Curr. Biol. 2012, 22, 590–595. [Google Scholar] [CrossRef] [Green Version]
- Perry, V.H.; Brown, M.C.; Lunn, E.R.; Tree, P.; Gordon, S. Evidence that Very Slow Wallerian Degeneration in C57BL/Ola Mice is an Intrinsic Property of the Peripheral Nerve. Eur. J. Neurosci. 1990, 2, 802–808. [Google Scholar] [CrossRef] [PubMed]
- Avery, M.A.; Rooney, T.M.; Pandya, J.D.; Wishart, T.M.; Gillingwater, T.H.; Geddes, J.W.; Sullivan, P.G.; Freeman, M.R. Wld S prevents axon degeneration through increased mitochondrial flux and enhanced mitochondrial Ca 2+ buffering. Curr. Biol. 2012, 22, 596–600. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Yap, C.C.; Winckler, B. Harnessing the Power of the Endosome to Regulate Neural Development. Neuron 2012, 74, 440–451. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Hausott, B.; Klimaschewski, L. Membrane turnover and receptor trafficking in regenerating axons. Eur. J. Neurosci. 2016, 43, 309–317. [Google Scholar] [CrossRef]
- Tojima, T.; Kamiguchi, H. Exocytic and endocytic membrane trafficking in axon development. Dev. Growth Differ. 2015, 57, 291–304. [Google Scholar] [CrossRef] [Green Version]
- Villarroel-Campos, D.; Bronfman, F.C.; Gonzalez-Billault, C. Rab GTPase signaling in neurite outgrowth and axon specification. Cytoskeleton 2016, 73, 498–507. [Google Scholar] [CrossRef]
- Donaldson, J.G.; Johnson, D.L.; Dutta, D. Rab and Arf G proteins in endosomal trafficking and cell surface homeostasis. Small GTPases 2016, 7, 247–251. [Google Scholar] [CrossRef] [Green Version]
- Raiborg, C.; Stenmark, H. The ESCRT machinery in endosomal sorting of ubiquitylated membrane proteins. Nature 2009, 458, 445–452. [Google Scholar] [CrossRef] [PubMed]
- Stenmark, H. Rab GTPases as coordinators of vesicle traffic. Nat. Rev. Mol. Cell Biol. 2009, 10, 513–525. [Google Scholar] [CrossRef] [PubMed]
- Welz, T.; Wellbourne-Wood, J.; Kerkhoff, E. Orchestration of cell surface proteins by Rab11. Trends Cell Biol. 2014, 24, 407–415. [Google Scholar] [CrossRef]
- Van Bergeijk, P.; Adrian, M.; Hoogenraad, C.C.; Kapitein, L.C. Optogenetic control of organelle transport and positioning. Nature 2015, 518, 111–114. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Eva, R.; Dassie, E.; Caswell, P.T.; Dick, G.; ffrench-Constant, C.; Norman, J.C.; Fawcett, J.W. Rab11 and Its Effector Rab Coupling Protein Contribute to the Trafficking of 1 Integrins during Axon Growth in Adult Dorsal Root Ganglion Neurons and PC12 Cells. J. Neurosci. 2010, 30, 11654–11669. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Shirane, M.; Nakayama, K.I. Protrudin Induces Neurite Formation by Directional Membrane Trafficking. Science 2006, 314, 818–821. [Google Scholar] [CrossRef] [PubMed]
- Fujita, A.; Koinuma, S.; Yasuda, S.; Nagai, H.; Kamiguchi, H.; Wada, N.; Nakamura, T. GTP hydrolysis of TC10 promotes neurite outgrowth through exocytic fusion of Rab11- and L1-containing vesicles by releasing exocyst component Exo70. PLoS ONE 2013, 8, e79689. [Google Scholar] [CrossRef] [PubMed]
- Takano, T.; Urushibara, T.; Yoshioka, N.; Saito, T.; Fukuda, M.; Tomomura, M.; Hisanaga, S.I. LMTK1 regulates dendritic formation by regulating movement of Rab11A-positive endosomes. Mol. Biol. Cell 2014, 25, 1755–1768. [Google Scholar] [CrossRef] [PubMed]
- Takano, T.; Tomomura, M.; Yoshioka, N.; Tsutsumi, K.; Terasawa, Y.; Saito, T.; Kawano, H.; Kamiguchi, H.; Fukuda, M.; Hisanaga, S.I. LMTK1/AATYK1 is a novel regulator of axonal outgrowth that acts via Rab11 in a Cdk5-dependent manner. J. Neurosci. 2012, 32, 6587–6599. [Google Scholar] [CrossRef]
- Hisanaga, S.I.; Wei, R.; Huo, A.; Tomomura, M. LMTK1, a Novel Modulator of Endosomal Trafficking in Neurons. Front. Mol. Neurosci. 2020, 13. [Google Scholar] [CrossRef]
- Koseki, H.; Donegá, M.; Lam, B.Y.; Petrova, V.; van Erp, S.; Yeo, G.S.; Kwok, J.C.; ffrench-Constant, C.; Eva, R.; Fawcett, J.W. Selective rab11 transport and the intrinsic regenerative ability of CNS axons. Elife 2017, 6, e26956. [Google Scholar] [CrossRef]
- Montagnac, G.; Sibarita, J.B.; Loubéry, S.; Daviet, L.; Romao, M.; Raposo, G.; Chavrier, P. ARF6 Interacts with JIP4 to Control a Motor Switch Mechanism Regulating Endosome Traffic in Cytokinesis. Curr. Biol. 2009, 19, 184–195. [Google Scholar] [CrossRef] [Green Version]
- Eva, R.; Crisp, S.; Marland, J.R.K.; Norman, J.C.; Kanamarlapudi, V.; ffrench-Constant, C.; Fawcett, J.W. ARF6 Directs Axon Transport and Traffic of Integrins and Regulates Axon Growth in Adult DRG Neurons. J. Neurosci. 2012, 32, 10352–10364. [Google Scholar] [CrossRef] [Green Version]
- Franssen, E.H.P.; Zhao, R.-R.; Koseki, H.; Kanamarlapudi, V.; Hoogenraad, C.C.; Eva, R.; Fawcett, J.W. Exclusion of Integrins from CNS Axons Is Regulated by Arf6 Activation and the AIS. J. Neurosci. 2015, 35, 8359–8375. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Eva, R.; Koseki, H.; Kanamarlapudi, V.; Fawcett, J.W. EFA6 regulates selective polarised transport and axon regeneration from the axon initial segment. J. Cell Sci. 2017, 130, 3663–3675. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Quiroga, S.; Bisbal, M.; Cáceres, A. Regulation of plasma membrane expansion during axon formation. Dev. Neurobiol. 2018, 78, 170–180. [Google Scholar] [CrossRef] [PubMed]
- Winckler, B.; Yap, C.C. Endocytosis and Endosomes at the Crossroads of Regulating Trafficking of Axon Outgrowth-Modifying Receptors. Traffic 2011, 12, 1099–1108. [Google Scholar] [CrossRef] [Green Version]
- Rozés-Salvador, V.; González-Billault, C.; Conde, C. The Recycling Endosome in Nerve Cell Development: One Rab to Rule Them All? Front. Cell Dev. Biol. 2020, 8, 1479. [Google Scholar] [CrossRef] [PubMed]
- Falk, J.; Konopacki, F.A.; Zivraj, K.H.; Holt, C.E. Rab5 and rab4 regulate axon elongation in the Xenopus visual system. J. Neurosci. 2014, 34, 373–391. [Google Scholar] [CrossRef] [Green Version]
- Ponomareva, O.Y.; Eliceiri, K.W.; Halloran, M.C. Charcot-Marie-Tooth 2b associated Rab7 mutations cause axon growth and guidance defects during vertebrate sensory neuron development. Neural Dev. 2016, 11, 2. [Google Scholar] [CrossRef] [Green Version]
- Nakazawa, H.; Sada, T.; Toriyama, M.; Tago, K.; Sugiura, T.; Fukuda, M.; Inagaki, N. Rab33a mediates anterograde vesicular transport for membrane exocytosis and axon outgrowth. J. Neurosci. 2012, 32, 12712–12725. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Villarroel-Campos, D.; Henríquez, D.R.; Bodaleo, F.J.; Oguchi, M.E.; Bronfman, F.C.; Fukuda, M.; Gonzalez-Billault, C. Rab35 functions in axon elongation are regulated by P53-related protein kinase in a mechanism that involves Rab35 protein degradation and the microtubule-associated protein 1B. J. Neurosci. 2016, 36, 7298–7313. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Deng, C.Y.; Lei, W.L.; Xu, X.H.; Ju, X.C.; Liu, Y.; Luo, Z.G. JIP1 mediates anterograde transport of Rab10 cargos during neuronal polarization. J. Neurosci. 2014, 34, 1710–1723. [Google Scholar] [CrossRef] [Green Version]
- Liu, Y.; Xu, X.H.; Chen, Q.; Wang, T.; Deng, C.Y.; Song, B.L.; Du, J.L.; Luo, Z.G. Myosin Vb controls biogenesis of post-Golgi Rab10 carriers during axon development. Nat. Commun. 2013, 4, 2005. [Google Scholar] [CrossRef] [Green Version]
- Xu, X.H.; Deng, C.Y.; Liu, Y.; He, M.; Peng, J.; Wang, T.; Yuan, L.; Zheng, Z.S.; Blackshear, P.J.; Luo, Z.G. MARCKS regulates membrane targeting of Rab10 vesicles to promote axon development. Cell Res. 2014, 24, 576–594. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Hervera, A.; De Virgiliis, F.; Palmisano, I.; Zhou, L.; Tantardini, E.; Kong, G.; Hutson, T.; Danzi, M.C.; Perry, R.B.T.; Santos, C.X.C.; et al. Reactive oxygen species regulate axonal regeneration through the release of exosomal NADPH oxidase 2 complexes into injured axons. Nat. Cell Biol. 2018, 20, 307–319. [Google Scholar] [CrossRef]
- Sekine, Y.; Lin-Moore, A.; Chenette, D.M.; Wang, X.; Jiang, Z.; Cafferty, W.B.; Hammarlund, M.; Strittmatter, S.M. Functional Genome-wide Screen Identifies Pathways Restricting Central Nervous System Axonal Regeneration. Cell Rep. 2018, 23, 415–428. [Google Scholar] [CrossRef] [Green Version]
- Xu, H.; Ren, D. Lysosomal physiology. Annu. Rev. Physiol. 2015, 77, 57–80. [Google Scholar] [CrossRef] [Green Version]
- Ferguson, S.M. Axonal transport and maturation of lysosomes. Curr. Opin. Neurobiol. 2018, 51, 45–51. [Google Scholar] [CrossRef]
- Inpanathan, S.; Botelho, R.J. The lysosome signaling platform: Adapting with the Times. Front. Cell Dev. Biol. 2019, 7, 113. [Google Scholar] [CrossRef] [Green Version]
- Ballabio, A.; Bonifacino, J.S. Lysosomes as dynamic regulators of cell and organismal homeostasis. Nat. Rev. Mol. Cell Biol. 2020, 21, 101–118. [Google Scholar] [CrossRef] [PubMed]
- Farías, G.G.; Guardia, C.M.; De Pace, R.; Britt, D.J.; Bonifacino, J.S. BORC/kinesin-1 ensemble drives polarized transport of lysosomes into the axon. Proc. Natl. Acad. Sci. USA 2017, 114, E2955–E2964. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kenific, C.M.; Wittmann, T.; Debnath, J. Autophagy in adhesion and migration. J. Cell Sci. 2016, 129, 3685–3693. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Stavoe, A.K.H.; Holzbaur, E.L.F. Autophagy in neurons. Annu. Rev. Cell Dev. Biol. 2019, 35, 477–500. [Google Scholar] [CrossRef]
- Maday, S.; Holzbaur, E.L.F. Compartment-Specific Regulation of Autophagy in Primary Neurons. J. Neurosci. 2016, 36, 5933–5945. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Maday, S.; Wallace, K.E.; Holzbaur, E.L.F. Autophagosomes initiate distally and mature during transport toward the cell soma in primary neurons. J. Cell Biol. 2012, 196, 407–417. [Google Scholar] [CrossRef]
- Maday, S.; Holzbaur, E.L.F. Article Autophagosome Biogenesis in Primary Neurons Follows an Ordered and Spatially Regulated Pathway. Dev. Cell 2014, 30, 71–85. [Google Scholar] [CrossRef] [Green Version]
- Stavoe, A.K.H.; Hill, S.E.; Hall, D.H.; Colón-Ramos, D.A. KIF1A/UNC-104 Transports ATG-9 to Regulate Neurodevelopment and Autophagy at Synapses. Dev. Cell 2016, 38, 171–185. [Google Scholar] [CrossRef] [Green Version]
- Puri, C.; Vicinanza, M.; Ashkenazi, A.; Gratian, M.J.; Zhang, Q.; Bento, C.F.; Renna, M.; Menzies, F.M.; Rubinsztein, D.C. The RAB11A-Positive Compartment Is a Primary Platform for Autophagosome Assembly Mediated by WIPI2 Recognition of PI3P-RAB11A. Dev. Cell 2018, 45, 114–131.e8. [Google Scholar] [CrossRef] [Green Version]
- Shibata, M.; Lu, T.; Furuya, T.; Degterev, A.; Mizushima, N.; Yoshimori, T.; MacDonald, M.; Yankner, B.; Yuan, J. Regulation of intracellular accumulation of mutant huntingtin by beclin 1. J. Biol. Chem. 2006, 281, 14474–14485. [Google Scholar] [CrossRef] [Green Version]
- Simonsen, A.; Cumming, R.C.; Brech, A.; Isakson, P.; Schubert, D.R.; Finley, K.D. Promoting basal levels of autophagy in the nervous system enhances longevity and oxidant resistance in adult Drosophila. Autophagy 2008, 4, 176–184. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Lipinski, M.M.; Zheng, B.; Lu, T.; Yan, Z.; Py, B.F.; Ng, A.; Xavier, R.J.; Li, C.; Yankner, B.A.; Scherzer, C.R.; et al. Genome-wide analysis reveals mechanisms modulating autophagy in normal brain aging and in Alzheimer’s disease. Proc. Natl. Acad. Sci. USA 2010, 107, 14164–14169. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Chang, J.T.; Kumsta, C.; Hellman, A.B.; Adams, L.M.; Hansen, M. Spatiotemporal regulation of autophagy during Caenorhabditis elegans aging. Elife 2017, 6, e18459. [Google Scholar] [CrossRef] [PubMed]
- Glatigny, M.; Moriceau, S.; Rivagorda, M.; Ramos-Brossier, M.; Nascimbeni, A.C.; Lante, F.; Shanley, M.R.; Boudarene, N.; Rousseaud, A.; Friedman, A.K.; et al. Autophagy Is Required for Memory Formation and Reverses Age-Related Memory Decline. Curr. Biol. 2019, 29, 435–448.e8. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Sakamoto, K.; Ozaki, T.; Ko, Y.C.; Tsai, C.F.; Gong, Y.; Morozumi, M.; Ishikawa, Y.; Uchimura, K.; Nadanaka, S.; Kitagawa, H.; et al. Glycan sulfation patterns define autophagy flux at axon tip via PTPRσ-cortactin axis. Nat. Chem. Biol. 2019, 15, 699–709. [Google Scholar] [CrossRef]
- Tran, A.P.; Warren, P.M.; Silver, J. Regulation of autophagy by inhibitory CSPG interactions with receptor PTPσ and its impact on plasticity and regeneration after spinal cord injury. Exp. Neurol. 2020, 328, 113276. [Google Scholar] [CrossRef]
- Galluzzi, L.; Green, D.R. Autophagy-Independent Functions of the Autophagy Machinery. Cell 2019, 177, 1682–1699. [Google Scholar] [CrossRef]
- He, M.; Ding, Y.; Chu, C.; Tang, J.; Xiao, Q.; Luo, Z.G. Autophagy induction stabilizes microtubules and promotes axon regeneration after spinal cord injury. Proc. Natl. Acad. Sci. USA 2016, 113, 11324–11329. [Google Scholar] [CrossRef] [Green Version]
- Kannan, M.; Bayam, E.; Wagner, C.; Rinaldi, B.; Kretz, P.F.; Tilly, P.; Roos, M.; McGillewie, L.; Bär, S.; Minocha, S.; et al. WD40-repeat 47, a microtubule-associated protein, is essential for brain development and autophagy. Proc. Natl. Acad. Sci. USA 2017, 114, E9308–E9317. [Google Scholar] [CrossRef] [Green Version]
- Wang, B.; Iyengar, R.; Li-Harms, X.; Joo, J.H.; Wright, C.; Lavado, A.; Horner, L.; Yang, M.; Guan, J.L.; Frase, S.; et al. The autophagy-inducing kinases, ULK1 and ULK2, regulate axon guidance in the developing mouse forebrain via a noncanonical pathway. Autophagy 2018, 14, 796–811. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Nikoletopoulou, V.; Sidiropoulou, K.; Kallergi, E.; Dalezios, Y.; Tavernarakis, N. Modulation of Autophagy by BDNF Underlies Synaptic Plasticity. Cell Metab. 2017, 26, 230–242.e5. [Google Scholar] [CrossRef] [Green Version]
- Ban, B.-K.; Jun, M.-H.; Ryu, H.-H.; Jang, D.-J.; Ahmad, S.T.; Lee, J.-A. Autophagy Negatively Regulates Early Axon Growth in Cortical Neurons. Mol. Cell. Biol. 2013, 33, 3907–3919. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Clarke, J.-P.; Mearow, K. Autophagy inhibition in endogenous and nutrient-deprived conditions reduces dorsal root ganglia neuron survival and neurite growth in vitro. J. Neurosci. Res. 2016, 94, 653–670. [Google Scholar] [CrossRef]
- Rabanal-Ruiz, Y.; Korolchuk, V.I. mTORC1 and nutrient homeostasis: The central role of the lysosome. Int. J. Mol. Sci. 2018, 19, 818. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Wei, X.; Luo, L.; Chen, J. Roles of mTOR Signaling in Tissue Regeneration. Cells 2019, 8, 1075. [Google Scholar] [CrossRef] [Green Version]
- Kim, J.; Guan, K.L. mTOR as a central hub of nutrient signalling and cell growth. Nat. Cell Biol. 2019, 21, 63–71. [Google Scholar] [CrossRef] [PubMed]
- Abe, N.; Borson, S.H.; Gambello, M.J.; Wang, F.; Cavalli, V. Mammalian Target of Rapamycin (mTOR) activation increases axonal growth capacity of injured peripheral nerves. J. Biol. Chem. 2010, 285, 28034–28043. [Google Scholar] [CrossRef] [Green Version]
- Danilov, C.A.; Steward, O. Conditional genetic deletion of PTEN after a spinal cord injury enhances regenerative growth of CST axons and motor function recovery in mice. Exp. Neurol. 2015, 266, 147–160. [Google Scholar] [CrossRef] [Green Version]
- Geoffroy, C.G.; Hilton, B.J.; Tetzlaff, W.; Zheng, B. Evidence for an Age-Dependent Decline in Axon Regeneration in the Adult Mammalian Central Nervous System. Cell Rep. 2016, 15, 238–246. [Google Scholar] [CrossRef] [Green Version]
- Leibinger, M.; Andreadaki, A.; Golla, R.; Levin, E.; Hilla, A.M.; Diekmann, H.; Fischer, D. Boosting CNS axon regeneration by harnessing antagonistic effects of GSK3 activity. Proc. Natl. Acad. Sci. USA 2017, 114, E5454–E5463. [Google Scholar] [CrossRef] [Green Version]
- Leibinger, M.; Hilla, A.M.; Andreadaki, A.; Fischer, D. GSK3-CRMP2 signaling mediates axonal regeneration induced by Pten knockout. Commun. Biol. 2019, 2, 318. [Google Scholar] [CrossRef] [Green Version]
- Miao, L.; Yang, L.; Huang, H.; Liang, F.; Ling, C.; Hu, Y. MTORC1 is necessary but mTORC2 and GSK3β are inhibitory for AKT3-induced axon regeneration in the central nervous system. Elife 2016, 5, e14908. [Google Scholar] [CrossRef] [PubMed]
- Tassew, N.G.; Charish, J.; Shabanzadeh, A.P.; Luga, V.; Harada, H.; Farhani, N.; D’Onofrio, P.; Choi, B.; Ellabban, A.; Nickerson, P.E.B.; et al. Exosomes Mediate Mobilization of Autocrine Wnt10b to Promote Axonal Regeneration in the Injured CNS. Cell Rep. 2017, 20, 99–111. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Pita-Thomas, W.; Mahar, M.; Joshi, A.; Gan, D.; Cavalli, V. HDAC5 promotes optic nerve regeneration by activating the mTOR pathway. Exp. Neurol. 2019, 317, 271–283. [Google Scholar] [CrossRef] [PubMed]
- Poulopoulos, A.; Murphy, A.J.; Ozkan, A.; Davis, P.; Hatch, J.; Kirchner, R.; Macklis, J.D. Subcellular transcriptomes and proteomes of developing axon projections in the cerebral cortex. Nature 2019, 565, 356–360. [Google Scholar] [CrossRef]
- Rodríguez, A.; Webster, P.; Ortego, J.; Andrews, N.W. Lysosomes behave as Ca2+-regulated exocytic vesicles in fibroblasts and epithelial cells. J. Cell Biol. 1997, 137, 93–104. [Google Scholar] [CrossRef]
- Reddy, A.; Caler, E.V.; Andrews, N.W. Plasma membrane repair is mediated by Ca2+-regulated exocytosis of lysosomes. Cell 2001, 106, 157–169. [Google Scholar] [CrossRef] [Green Version]
- Jaiswal, J.K.; Andrews, N.W.; Simon, S.M. Membrane proximal lysosomes are the major vesicles responsible for calcium-dependent exocytosis in nonsecretory cells. J. Cell Biol. 2002, 159, 625–635. [Google Scholar] [CrossRef] [Green Version]
- Arantes, R.M.E.; Andrews, N.W. A role for synaptotagmin VII-regulated exocytosis of lysosomes in neurite outgrowth from primary sympathetic neurons. J. Neurosci. 2006, 26, 4630–4637. [Google Scholar] [CrossRef] [Green Version]
- Czibener, C.; Sherer, N.M.; Becker, S.M.; Pypaert, M.; Hui, E.; Chapman, E.R.; Mothes, W.; Andrews, N.W. Ca2+ and synaptotagmin VII-dependent delivery of lysosomal membrane to nascent phagosomes. J. Cell Biol. 2006, 174, 997–1007. [Google Scholar] [CrossRef] [Green Version]
- Sato, M.; Yoshimura, S.; Hirai, R.; Goto, A.; Kunii, M.; Atik, N.; Sato, T.; Sato, K.; Harada, R.; Shimada, J.; et al. The role of VAMP7/TI-VAMP in cell polarity and lysosomal exocytosis in vivo. Traffic 2011, 12, 1383–1393. [Google Scholar] [CrossRef]
- Naegeli, K.M.; Hastie, E.; Garde, A.; Wang, Z.; Keeley, D.P.; Gordon, K.L.; Pani, A.M.; Kelley, L.C.; Morrissey, M.A.; Chi, Q.; et al. Cell Invasion In Vivo via Rapid Exocytosis of a Transient Lysosome-Derived Membrane Domain. Dev. Cell 2017, 43, 403–417.e10. [Google Scholar] [CrossRef] [Green Version]
- Jiang, M.; Meng, J.; Zeng, F.; Qing, H.; Hook, G.; Hook, V.; Wu, Z.; Ni, J. Cathepsin B inhibition blocks neurite outgrowth in cultured neurons by regulating lysosomal trafficking and remodeling. J. Neurochem. 2020, 155, 300–312. [Google Scholar] [CrossRef]
- Jung, J.; Shin, Y.H.; Konishi, H.; Lee, S.J.; Kiyama, H. Possible ATP release through lysosomal exocytosis from primary sensory neurons. Biochem. Biophys. Res. Commun. 2013, 430, 488–493. [Google Scholar] [CrossRef]
- Tam, C.; Idone, V.; Devlin, C.; Fernandes, M.C.; Flannery, A.; He, X.; Schuchman, E.; Tabas, I.; Andrews, N.W. Exocytosis of acid sphingomyelinase by wounded cells promotes endocytosis and plasma membrane repair. J. Cell Biol. 2010, 189, 1027–1038. [Google Scholar] [CrossRef] [Green Version]
- Castro-Gomes, T.; Corrotte, M.; Tam, C.; Andrews, N.W. Plasma membrane repair is regulated extracellularly by proteases released from lysosomes. PLoS ONE 2016, 11, e0152583. [Google Scholar]
- Korhonen, L.; Lindholm, D. The ubiquitin proteasome system in synaptic and axonal degeneration: A new twist to an old cycle. J. Cell Biol. 2004, 165, 27–30. [Google Scholar] [CrossRef] [PubMed]
- Lee, M.; Liu, Y.C.; Chen, C.; Lu, C.H.; Lu, S.T.; Huang, T.N.; Hsu, M.T.; Hsueh, Y.P.; Cheng, P.L. Ecm29-mediated proteasomal distribution modulates excitatory GABA responses in the developing brain. J. Cell Biol. 2020, 219, e201903033. [Google Scholar] [CrossRef] [PubMed]
- Minis, A.; Rodriguez, J.A.; Levin, A.; Liu, K.; Govek, E.E.; Hatten, M.E.; Steller, H. The proteasome regulator PI31 is required for protein homeostasis, synapse maintenance, and neuronal survival in mice. Proc. Natl. Acad. Sci. USA 2019, 116, 24639–24650. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Otero, M.G.; Alloatti, M.; Cromberg, L.E.; Almenar-Queralt, A.; Encalada, S.E.; Pozo Devoto, V.M.; Bruno, L.; Goldstein, L.S.B.; Falzone, T.L. Fast axonal transport of the proteasome complex depends on membrane interaction and molecular motor function. J. Cell Sci. 2014, 127, 1537–1549. [Google Scholar] [CrossRef] [Green Version]
- Bingol, B.; Schuman, E.M. Activity-dependent dynamics and sequestration of proteasomes in dendritic spines. Nature 2006, 441, 1144–1148. [Google Scholar] [CrossRef] [Green Version]
- Hamilton, A.M.; Zito, K. Breaking it down: The ubiquitin proteasome system in neuronal morphogenesis. Neural Plast. 2013, 2013, 196848. [Google Scholar] [CrossRef] [PubMed]
- DiAntonio, A.; Hicke, L. Ubiquitin-Dependent Regulation of the Synapse. Annu. Rev. Neurosci. 2004, 27, 223–246. [Google Scholar] [CrossRef] [PubMed]
- Ohtani-Kaneko, R.; Takada, K.; Iigo, M.; Hara, M.; Yokosawa, H.; Kawashima, S.; Ohkawa, K.; Hirata, K. Proteasome inhibitors which induce neurite outgrowth from PC12h cells cause different subcellular accumulations of multi-ubiquitin chains. Neurochem. Res. 1998, 23, 1435–1443. [Google Scholar] [CrossRef] [PubMed]
- Obin, M.; Mesco, E.; Gong, X.; Haas, A.L.; Joseph, J.; Taylor, A. Neurite outgrowth in PC12 cells: Distinguishing the roles of ubiquitylation and ubiquitin-dependent proteolysis. J. Biol. Chem. 1999, 274, 11789–11795. [Google Scholar] [CrossRef] [Green Version]
- Ōmura, S.; Fujimoto, T.; Otoguro, K.; Matsuzaki, K.; Moriguchi, R.; Tanaka, H.; Sasaki, Y. Lactacystin, a novel microbial metabolite, induces neurito-genesis of neuroblastoma cells. J. Antibiot. 1991, 44, 113–116. [Google Scholar] [CrossRef] [Green Version]
- Klimaschewski, L.; Hausott, B.; Ingorokva, S.; Pfaller, K. Constitutively expressed catalytic proteasomal subunits are up-regulated during neuronal differentiation and required for axon initiation, elongation and maintenance. J. Neurochem. 2006, 96, 1708–1717. [Google Scholar] [CrossRef]
- Laser, H.; Mack, T.G.A.; Wagner, D.; Coleman, M.P. Proteasome inhibition arrests neurite outgrowth and causes dying-back degeneration in primary culture. J. Neurosci. Res. 2003, 74, 906–916. [Google Scholar] [CrossRef]
- Zheng, Q.; Huang, T.; Zhang, L.; Zhou, Y.; Luo, H.; Xu, H.; Wang, X. Dysregulation of ubiquitin-proteasome system in neurodegenerative diseases. Front. Aging Neurosci. 2016, 8, 303. [Google Scholar] [CrossRef] [PubMed]
- Njomen, E.; Tepe, J.J. Proteasome Activation as a New Therapeutic Approach to Target Proteotoxic Disorders. J. Med. Chem. 2019, 62, 6469–6481. [Google Scholar] [CrossRef]
- Tanaka, K.; Matsuda, N. Proteostasis and neurodegeneration: The roles of proteasomal degradation and autophagy. Biochim. Biophys. Acta-Mol. Cell Res. 2014, 1843, 197–204. [Google Scholar] [CrossRef] [Green Version]
- Jin, E.J.; Ko, H.R.; Hwang, I.; Kim, B.S.; Choi, J.Y.; Park, K.W.; Cho, S.W.; Ahn, J.Y. Akt regulates neurite growth by phosphorylation-dependent inhibition of radixin proteasomal degradation. Sci. Rep. 2018, 8, 2557. [Google Scholar] [CrossRef] [PubMed]
- Tursun, B.; Schlüter, A.; Peters, M.A.; Viehweger, B.; Ostendorff, H.P.; Soosairajah, J.; Drung, A.; Bossenz, M.; Johnsen, S.A.; Schweizer, M.; et al. The ubiquitin ligase Rnf6 regulates local LIM kinase 1 levels in axonal growth cones. Genes Dev. 2005, 19, 2307–2319. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Yan, D.; Guo, L.; Wang, Y. Requirement of dendritic Akt degradation by the ubiquitin-proteasome system for neuronal polarity. J. Cell Biol. 2006, 174, 415–424. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Takabatake, M.; Goshima, Y.; Sasaki, Y. Semaphorin-3A Promotes Degradation of Fragile X Mental Retardation Protein in Growth Cones via the Ubiquitin-Proteasome Pathway. Front. Neural Circuits 2020, 14, 5. [Google Scholar] [CrossRef]
- Campbell, D.S.; Holt, C.E. Chemotropic responses of retinal growth cones mediated by rapid local protein synthesis and degradation. Neuron 2001, 32, 1013–1026. [Google Scholar] [CrossRef] [Green Version]
- Hsu, M.-T.; Guo, C.-L.; Liou, A.Y.; Chang, T.-Y.; Ng, M.-C.; Florea, B.I.; Overkleeft, H.S.; Wu, Y.-L.; Liao, J.-C.; Cheng, P.-L. Stage-Dependent Axon Transport of Proteasomes Contributes to Axon Development. Dev. Cell 2015, 35, 418–431. [Google Scholar] [CrossRef] [Green Version]
- Knöferle, J.; Ramljak, S.; Koch, J.C.; Tönges, L.; Asif, A.R.; Michel, U.; Wouters, F.S.; Heermann, S.; Krieglstein, K.; Zerr, I.; et al. TGF-β 1 enhances neurite outgrowth via regulation of proteasome function and EFABP. Neurobiol. Dis. 2010, 38, 395–404. [Google Scholar] [CrossRef]
- Park, J.Y.; Jang, S.Y.; Shin, Y.K.; Suh, D.J.; Park, H.T. Calcium-dependent proteasome activation is required for axonal neurofilament degradation. Neural Regen. Res. 2013, 8, 3401. [Google Scholar]
- Staal, J.A.; Dickson, T.C.; Chung, R.S.; Vickers, J.C. Disruption of the Ubiquitin Proteasome System following Axonal Stretch Injury Accelerates Progression to Secondary Axotomy. J. Neurotrauma 2009, 26, 781–788. [Google Scholar] [CrossRef]
- Valm, A.M.; Cohen, S.; Legant, W.R.; Melunis, J.; Hershberg, U.; Wait, E.; Cohen, A.R.; Davidson, M.W.; Betzig, E.; Lippincott-Schwartz, J. Applying systems-level spectral imaging and analysis to reveal the organelle interactome. Nature 2017, 546, 162–167. [Google Scholar] [CrossRef]
- Lee, S.; Wang, W.; Hwang, J.; Namgung, U.; Min, K.T. Increased ER–mitochondria tethering promotes axon regeneration. Proc. Natl. Acad. Sci. USA 2019, 116, 16074–16079. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Krauß, M.; Haucke, V. A grab to move on: ER –endosome contacts in membrane protrusion formation and neurite outgrowth. EMBO J. 2015, 34, 1442–1444. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Wu, H.; Carvalho, P.; Voeltz, G.K. Here, there, and everywhere: The importance of ER membrane contact sites. Science 2018, 361. [Google Scholar] [CrossRef] [Green Version]
- Mannan, A.U.; Krawen, P.; Sauter, S.M.; Boehm, J.; Chronowska, A.; Paulus, W.; Neesen, J.; Engel, W. ZFYVE27 (SPG33), a Novel Spastin-Binding Protein, Is Mutated in Hereditary Spastic Paraplegia. Am. J. Hum. Genet. 2006, 79, 351–357. [Google Scholar] [CrossRef] [Green Version]
- Evans, K.; Keller, C.; Pavur, K.; Glasgow, K.; Conn, B.; Lauring, B. Interaction of two hereditary spastic paraplegia gene products, spastin and atlastin, suggests a common pathway for axonal maintenance. Proc. Natl. Acad. Sci. USA 2006, 103, 10666–10671. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Züchner, S.; Wang, G.; Tran-Viet, K.N.; Nance, M.A.; Gaskell, P.C.; Vance, J.M.; Ashley-Koch, A.E.; Pericak-Vance, M.A. Mutations in the novel mitochondrial protein REEP1 cause hereditary spastic paraplegia type 31. Am. J. Hum. Genet. 2006, 79, 365–369. [Google Scholar] [CrossRef] [Green Version]
- Beetz, C.; Schüle, R.; Deconinck, T.; Tran-Viet, K.N.; Zhu, H.; Kremer, B.P.H.; Frints, S.G.M.; Van Zelst-Stams, W.A.G.; Byrne, P.; Otto, S.; et al. REEP1 mutation spectrum and genotype/phenotype correlation in hereditary spastic paraplegia type 31. Brain 2008, 131, 1078–1086. [Google Scholar] [CrossRef] [Green Version]
- Nishimura, A.L.; Mitne-Neto, M.; Silva, H.C.A.; Richieri-Costa, A.; Middleton, S.; Cascio, D.; Kok, F.; Oliveira, J.R.M.; Gillingwater, T.; Webb, J.; et al. A mutation in the vesicle-trafficking protein VAPB causes late-onset spinal muscular atrophy and amyotrophic lateral sclerosis. Am. J. Hum. Genet. 2004, 75, 822–831. [Google Scholar] [CrossRef] [Green Version]
- Kitada, T.; Asakawa, S.; Hattori, N.; Matsumine, H.; Yamamura, Y.; Minoshima, S.; Yokochi, M.; Mizuno, Y.; Shimizu, N. Mutations in the parkin gene cause autosomal recessive juvenile parkinsonism. Nature 1998, 392, 605–608. [Google Scholar] [CrossRef] [PubMed]
- Valente, E.M.; Abou-Sleiman, P.M.; Caputo, V.; Muqit, M.M.K.; Harvey, K.; Gispert, S.; Ali, Z.; Del Turco, D.; Bentivoglio, A.R.; Healy, D.G.; et al. Hereditary early-onset Parkinson’s disease caused by mutations in PINK1. Science 2004, 304, 1158–1160. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Valente, E.M.; Salvi, S.; Ialongo, T.; Marongiu, R.; Elia, A.E.; Caputo, V.; Romito, L.; Albanese, A.; Dallapiccola, B.; Bentivoglio, A.R. PINK1 mutations are associated with sporadic early-onset Parkinsonism. Ann. Neurol. 2004, 56, 336–341. [Google Scholar] [CrossRef] [PubMed]
- Züchner, S.; Mersiyanova, I.V.; Muglia, M.; Bissar-Tadmouri, N.; Rochelle, J.; Dadali, E.L.; Zappia, M.; Nelis, E.; Patitucci, A.; Senderek, J.; et al. Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat. Genet. 2004, 36, 449–451. [Google Scholar] [CrossRef] [PubMed]
- Sidiropoulos, P.N.M.; Miehe, M.; Bock, T.; Tinelli, E.; Oertli, C.I.; Kuner, R.; Meijer, D.; Wollscheid, B.; Niemann, A.; Suter, U. Dynamin 2 mutations in Charcot-Marie-Tooth neuropathy highlight the importance of clathrin-mediated endocytosis in myelination. Brain 2012, 135, 1395–1411. [Google Scholar] [CrossRef] [Green Version]
- Böhm, J.; Bulla, M.; Urquhart, J.E.; Malfatti, E.; Williams, S.G.; O’Sullivan, J.; Szlauer, A.; Koch, C.; Baranello, G.; Mora, M.; et al. ORAI1 Mutations with Distinct Channel Gating Defects in Tubular Aggregate Myopathy. Hum. Mutat. 2017, 38, 426–438. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Xu, L.; Wang, X.; Tong, C. Endoplasmic Reticulum–Mitochondria Contact Sites and Neurodegeneration. Front. Cell Dev. Biol. 2020, 8, 428. [Google Scholar] [CrossRef]
- Rieusset, J. The role of endoplasmic reticulum-mitochondria contact sites in the control of glucose homeostasis: An update. Cell Death Dis. 2018, 9, 388. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Rowland, A.A.; Voeltz, G.K. Endoplasmic reticulum-mitochondria contacts: Function of the junction. Nat. Rev. Mol. Cell Biol. 2012, 13, 607–615. [Google Scholar] [CrossRef] [Green Version]
- Marchi, S.; Patergnani, S.; Pinton, P. The endoplasmic reticulum-mitochondria connection: One touch, multiple functions. Biochim. Biophys. Acta-Bioenerg. 2014, 1837, 461–469. [Google Scholar] [CrossRef] [Green Version]
- Bravo-Sagua, R.; Torrealba, N.; Paredes, F.; Morales, P.E.; Pennanen, C.; López-Crisosto, C.; Troncoso, R.; Criollo, A.; Chiong, M.; Hill, J.A.; et al. Organelle communication: Signaling crossroads between homeostasis and disease. Int. J. Biochem. Cell Biol. 2014, 50, 55–59. [Google Scholar] [CrossRef]
- Grimm, S. The ER-mitochondria interface: The social network of cell death. Biochim. Biophys. Acta-Mol. Cell Res. 2012, 1823, 327–334. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Bernard-Marissal, N.; Médard, J.J.; Azzedine, H.; Chrast, R. Dysfunction in endoplasmic reticulum-mitochondria crosstalk underlies SIGMAR1 loss of function mediated motor neuron degeneration. Brain 2015, 138, 875–890. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Bernard-Marissal, N.; Chrast, R.; Schneider, B.L. Endoplasmic reticulum and mitochondria in diseases of motor and sensory neurons: A broken relationship? Cell Death Dis. 2018, 9, 1–16. [Google Scholar] [CrossRef]
- Stoica, R.; De Vos, K.J.; Paillusson, S.; Mueller, S.; Sancho, R.M.; Lau, K.F.; Vizcay-Barrena, G.; Lin, W.L.; Xu, Y.F.; Lewis, J.; et al. ER-mitochondria associations are regulated by the VAPB-PTPIP51 interaction and are disrupted by ALS/FTD-associated TDP-43. Nat. Commun. 2014, 5, 399. [Google Scholar] [CrossRef] [Green Version]
- Guardia-Laguarta, C.; Area-Gomez, E.; Rüb, C.; Liu, Y.; Magrané, J.; Becker, D.; Voos, W.; Schon, E.A.; Przedborski, S. α-synuclein is localized to mitochondria-associated ER membranes. J. Neurosci. 2014, 34, 249–259. [Google Scholar] [CrossRef]
- Schon, E.A.; Area-Gomez, E. Mitochondria-associated ER membranes in Alzheimer disease. Mol. Cell. Neurosci. 2013, 55, 26–36. [Google Scholar] [CrossRef]
- Calì, T.; Ottolini, D.; Negro, A.; Brini, M. α-synuclein controls mitochondrial calcium homeostasis by enhancing endoplasmic reticulum-mitochondria interactions. J. Biol. Chem. 2012, 287, 17914–17929. [Google Scholar] [CrossRef] [Green Version]
- Zampese, E.; Fasolato, C.; Kipanyula, M.J.; Bortolozzi, M.; Pozzan, T.; Pizzo, P. Presenilin 2 modulates endoplasmic reticulum (ER)-mitochondria interactions and Ca2+ cross-talk. Proc. Natl. Acad. Sci. USA 2011, 108, 2777–2782. [Google Scholar] [CrossRef] [Green Version]
- Szabadkai, G.; Bianchi, K.; Várnai, P.; De Stefani, D.; Wieckowski, M.R.; Cavagna, D.; Nagy, A.I.; Balla, T.; Rizzuto, R. Chaperone-mediated coupling of endoplasmic reticulum and mitochondrial Ca2+ channels. J. Cell Biol. 2006, 175, 901–911. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Trigo, D.; Goncalves, M.B.; Corcoran, J.P.T. The regulation of mitochondrial dynamics in neurite outgrowth by retinoic acid receptor β signaling. FASEB J. 2019, 33, 7225–7235. [Google Scholar] [CrossRef] [Green Version]
- Friedman, J.R.; DiBenedetto, J.R.; West, M.; Rowland, A.A.; Voeltz, G.K. Endoplasmic reticulum-endosome contact increases as endosomes traffic and mature. Mol. Biol. Cell 2013, 24, 1030–1040. [Google Scholar] [CrossRef] [PubMed]
- Raiborg, C.; Wenzel, E.M.; Stenmark, H. ER –endosome contact sites: Molecular compositions and functions. EMBO J. 2015, 34, 1848–1858. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Lu, M.; van Tartwijk, F.W.; Lin, J.Q.; Nijenhuis, W.; Parutto, P.; Fantham, M.; Christensen, C.N.; Avezov, E.; Holt, C.E.; Tunnacliffe, A.; et al. The structure and global distribution of the endoplasmic reticulum network are actively regulated by lysosomes. Sci. Adv. 2020, 6, eabc7209. [Google Scholar] [CrossRef]
- Raiborg, C.; Wenzel, E.M.; Pedersen, N.M.; Olsvik, H.; Schink, K.O.; Schultz, S.W.; Vietri, M.; Nisi, V.; Bucci, C.; Brech, A.; et al. Repeated ER–endosome contacts promote endosome translocation and neurite outgrowth. Nature 2015, 520, 234–238. [Google Scholar] [CrossRef] [PubMed]
- Allison, R.; Lumb, J.H.; Fassier, C.; Connell, J.W.; Martin, D.T.; Seaman, M.N.J.; Hazan, J.; Reid, E. An ESCRT-spastin interaction promotes fission of recycling tubules from the endosome. J. Cell Biol. 2013, 202, 527–543. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Allison, R.; Edgar, J.R.; Pearson, G.; Rizo, T.; Newton, T.; Günther, S.; Berner, F.; Hague, J.; Connell, J.W.; Winkler, J.; et al. Defects in ER-endosome contacts impact lysosome function in hereditary spastic paraplegia. J. Cell Biol. 2017, 216, 1337–1355. [Google Scholar] [CrossRef] [PubMed]
- Eden, E.R.; White, I.J.; Tsapara, A.; Futter, C.E. Membrane contacts between endosomes and ER provide sites for PTP1B-epidermal growth factor receptor interaction. Nat. Cell Biol. 2010, 12, 267–272. [Google Scholar] [CrossRef]
- Eden, E.R.; Sanchez-Heras, E.; Tsapara, A.; Sobota, A.; Levine, T.P.; Futter, C.E. Annexin A1 Tethers Membrane Contact Sites that Mediate ER to Endosome Cholesterol Transport. Dev. Cell 2016, 37, 473–483. [Google Scholar] [CrossRef] [Green Version]
- Kilpatrick, B.S.; Eden, E.R.; Hockey, L.N.; Yates, E.; Futter, C.E.; Patel, S. An Endosomal NAADP-Sensitive Two-Pore Ca2+ Channel Regulates ER-Endosome Membrane Contact Sites to Control Growth Factor Signaling. Cell Rep. 2017, 18, 1636–1645. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Petkovic, M.; Oses-Prieto, J.; Burlingame, A.; Jan, L.Y.; Jan, Y.N. TMEM16K is an interorganelle regulator of endosomal sorting. Nat. Commun. 2020, 11, 1–16. [Google Scholar] [CrossRef] [PubMed]
- Elbaz-Alon, Y.; Guo, Y.; Segev, N.; Harel, M.; Quinnell, D.E.; Geiger, T.; Avinoam, O.; Li, D.; Nunnari, J. PDZD8 interacts with Protrudin and Rab7 at ER-late endosome membrane contact sites associated with mitochondria. Nat. Commun. 2020, 11, 1–14. [Google Scholar] [CrossRef] [PubMed]
- Hirabayashi, Y.; Kwon, S.K.; Paek, H.; Pernice, W.M.; Paul, M.A.; Lee, J.; Erfani, P.; Raczkowski, A.; Petrey, D.S.; Pon, L.A.; et al. ER-mitochondria tethering by PDZD8 regulates Ca2+ dynamics in mammalian neurons. Science 2017, 358, 623–630. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Liao, Y.C.; Fernandopulle, M.S.; Wang, G.; Choi, H.; Hao, L.; Drerup, C.M.; Patel, R.; Qamar, S.; Nixon-Abell, J.; Shen, Y.; et al. RNA Granules Hitchhike on Lysosomes for Long-Distance Transport, Using Annexin A11 as a Molecular Tether. Cell 2019, 179, 147–164.e20. [Google Scholar] [CrossRef] [Green Version]
- Lee, J.E.; Cathey, P.I.; Wu, H.; Parker, R.; Voeltz, G.K. Endoplasmic reticulum contact sites regulate the dynamics of membraneless organelles. Science 2020, 367, eaay7108. [Google Scholar] [CrossRef] [PubMed]
- 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. 2012, 22, 545–552. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Miller, K.E.; Sheetz, M.P. Axonal mitochondrial transport and potential are correlated. J. Cell Sci. 2004, 117, 2791–2804. [Google Scholar] [CrossRef] [Green Version]
- Lin, M.Y.; Cheng, X.T.; Tammineni, P.; Xie, Y.; Zhou, B.; Cai, Q.; Sheng, Z.H. Releasing Syntaphilin Removes Stressed Mitochondria from Axons Independent of Mitophagy under Pathophysiological Conditions. Neuron 2017, 94, 595–610.e6. [Google Scholar] [CrossRef] [Green Version]
- Zheng, Y.; Zhang, X.; Wu, X.; Jiang, L.; Ahsan, A.; Ma, S.; Xiao, Z.; Han, F.; Qin, Z.H.; Hu, W.; et al. Somatic autophagy of axonal mitochondria in ischemic neurons. J. Cell Biol. 2019, 218, 1891–1907. [Google Scholar] [CrossRef] [Green Version]
- Wang, X.; Winter, D.; Ashrafi, G.; Schlehe, J.; Wong, Y.L.; Selkoe, D.; Rice, S.; Steen, J.; Lavoie, M.J.; Schwarz, T.L. PINK1 and Parkin target miro for phosphorylation and degradation to arrest mitochondrial motility. Cell 2011, 147, 893–906. [Google Scholar] [CrossRef] [Green Version]
- 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. 2014, 206, 655–670. [Google Scholar] [CrossRef] [PubMed]
- Gabriela Otero, M.; Fernandez Bessone, I.; Earle Hallberg, A.; Eneas Cromberg, L.; Cecilia De Rossi, M.; Saez, T.M.; Levi, V.; Almenar-Queralt, A.; Luis Falzone, T. Proteasome Stress Leads to APP Axonal Transport Defects by Promoting Its Amyloidogenic Processing in Lysosomes. 2018. Available online: http://jcs.biologists.org/content/joces/early/2018/05/02/jcs.214536.full.pdf (accessed on 6 March 2019).
Method for Manipulation of Autophagy | Neuronal Type Examined | Species and Age | Main Findings Regarding Neuronal Growth | Reference |
---|---|---|---|---|
Knockdown of ATG7 Application of 3-methyladenine | Primary cortical neurons | Embryonic rats and cultured for 1 till 3 days | Inhibition of autophagy resulted in: - elongation of neurites in vitro - reduction of RhoA signalling | Ban et al., 2013 [203] |
Knockout of: ATG-2, ATG9, ATG13, EPG-8, IGG-1, UNC104 | PVD nociceptive sensory neuron | Larval and adult C. elegans | Inhibition of autophagy resulted in: - elongation of the axon in vivo | Stavoe et al., 2016 [189] |
Knockout of: WDR47 | - Primary cortical and hippocampal neurons - Callosal and corticofugal neurons | - Embryonic mice and cultured for 4 days - Adult mice (16 weeks old) | Activation of autophagy resulted in: - Impaired formation and dynamics of growth cones in vitro -Defective and reduction of axonal projections in the corpus calossum in vivo - Destabilisation of microtubules | Kannan et al., 2017 [200] |
Knockout of: ATG-2, ATG9, ATG13, EPG-8, IGG-1, UNC104 | - HSN serotonergic motor neuron - DA9 cholinergic motor neuron - RIA interneuron - RIB interneuron - NSM pharyngeal neurosecretory-motor neuron | Larval and adult C. elegans | Inhibition of autophagy resulted in: - no phenotype in vivo | Stavoe et al., 2016 [189] |
Application of Tat-beclin1 | - Primary cortical neurons - Neuron types with axon fibres in the spinal cord | - Embryonic rats and cultured for 1 till 3 days - Adult mice (8 till 10 weeks old) subjected to spinal cord hemisection injury | Activation of autophagy resulted in: - enhanced neurite outgrowth on inhibitory substrates in vitro - Inhibition of axonal retraction after injury in cortical neurons in vitro and corticospinal neurons in vivo - Stimulation of axonal regeneration of monoaminergic neurons after injury in vivo - Stabilisation of microtubules | He et al., 2016 [199] |
Application of 3-methyladenine | Dorsal root ganglion neurons | Adult rat (4 till 5 weeks old) and cultured for 1 day | Inhibition of autophagy resulted in: - Reduction of neuronal survival - Inhibition of neurite growth and branching | Clarke and Mearow, 2016 [204] |
Membrane Contact Site | Disease | Genes Involved | Gene Function | References |
---|---|---|---|---|
ER- Endosomes/ ER-PM/ER-Lysosome | Hereditary Spastic Paraplegia | Zfyve27 | Vesicular membrane trafficking, ER-endosome/lysosome tethering | Mannan et al., 2006 [256] |
ER-Endosome | Hereditary Spastic Paraplegia | Spastin | Microtubule-severing protein | Evans et al., 2006 [257] |
ER-Mitochondria | Hereditary Spastic Paraplegia | REEP1 | Microtubule-mitochondria | Zuchner et al., 2006; Beetz et al., 2008 [258,259] |
ER-Endosome/ER-PM | Amyotrophic Lateral Sclerosis | VAP-A, VAP-B | ER-organelle tethering, facilitate protein interaction, vesicular trafficking | Nishimura et al., 2004 [260] |
ER-Mitochondria | Early onset autosomal Parkinson’s disease | PARKIN, PINK1 | Mitochondrial quality control and turnover | Kitada et al., 1998; Valente et al., 2004a; Valente et al., 2004b [261,262,263] |
ER-Mitochondria | Charcot Marie Tooth Disease | MFN-2 | Mitochondrial fusion, interaction with endoplasmic reticulum | Zuchner et al., 2004 [264] |
ER-Mitochondria | Charcot Marie Tooth Disease | DNM-2 | Vesicle trafficking, cytoskeleton dynamics, endosomal pathways | Sidiropoulos et al., 2012 [265] |
ER-PM | Tubular Aggregate Myopathy | STIM-1 | Calcium sensor | Bohm et al., 2017 [266] |
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Petrova, V.; Nieuwenhuis, B.; Fawcett, J.W.; Eva, R. Axonal Organelles as Molecular Platforms for Axon Growth and Regeneration after Injury. Int. J. Mol. Sci. 2021, 22, 1798. https://doi.org/10.3390/ijms22041798
Petrova V, Nieuwenhuis B, Fawcett JW, Eva R. Axonal Organelles as Molecular Platforms for Axon Growth and Regeneration after Injury. International Journal of Molecular Sciences. 2021; 22(4):1798. https://doi.org/10.3390/ijms22041798
Chicago/Turabian StylePetrova, Veselina, Bart Nieuwenhuis, James W. Fawcett, and Richard Eva. 2021. "Axonal Organelles as Molecular Platforms for Axon Growth and Regeneration after Injury" International Journal of Molecular Sciences 22, no. 4: 1798. https://doi.org/10.3390/ijms22041798