In situ structural mechanism of epothilone
Curcio, M. & Bradke, F. Axon regeneration in the central nervous system: facing the challenges from the inside. Annu. Rev. Cell Dev. Biol. 34, 495–521 (2018).
Google Scholar
He, Z. & Jin, Y. Intrinsic control of axon regeneration. Neuron 90, 437–451 (2016).
Google Scholar
Fawcett, J. W. The struggle to make CNS axons regenerate: why has it been so difficult? Neurochem. Res. 45, 144–158 (2020).
Google Scholar
Sharp, D. J., Scott, G. & Leech, R. Network dysfunction after traumatic brain injury. Nat. Rev. Neurol. 10, 156–166 (2014).
Google Scholar
Bradke, F., Fawcett, J. W. & Spira, M. E. Assembly of a new growth cone after axotomy: the precursor to axon regeneration. Nat. Rev. Neurosci. 13, 183–193 (2012).
Google Scholar
Schlaepfer, W. W. Calcium-induced degeneration of axoplasm in isolated segments of rat peripheral nerve. Brain Res. 69, 203–215 (1974).
Google Scholar
Wolf, J. A., Stys, P. K., Lusardi, T., Meaney, D. & Smith, D. H. Traumatic axonal injury induces calcium influx modulated by tetrodotoxin-sensitive sodium channels. J. Neurosci. 21, 1923–1930 (2001).
Google Scholar
Ziv, N. E. & Spira, M. E. Axotomy induces a transient and localized elevation of the free intracellular calcium concentration to the millimolar range. J. Neurophysiol. 74, 2625–2637 (1995).
Google Scholar
Schlaepfer, W. W. & Bunge, R. P. Effects of calcium ion concentration on the degeneration of amputated axons in tissue culture. J. Cell Biol. 59, 456–470 (1973).
Google Scholar
Ramon y Cajal, S. & May, R. M. Degeneration and Regeneration of the Nervous System (Oxford Univ. Press, 1928).
Li, D., Field, P. M. & Raisman, G. Failure of axon regeneration in postnatal rat entorhinohippocampal slice coculture is due to maturation of the axon, not that of the pathway or target. Eur. J. Neurosci. 7, 1164–1171 (1995).
Google Scholar
Tang-Schomer, M. D., Patel, A. R., Baas, P. W. & Smith, D. H. Mechanical breaking of microtubules in axons during dynamic stretch injury underlies delayed elasticity, microtubule disassembly, and axon degeneration. FASEB J. 24, 1401–1410 (2010).
Google Scholar
Erturk, A., Hellal, F., Enes, J. & Bradke, F. Disorganized microtubules underlie the formation of retraction bulbs and the failure of axonal regeneration. J. Neurosci. 27, 9169–9180 (2007).
Google Scholar
Blanquie, O. & Bradke, F. Cytoskeleton dynamics in axon regeneration. Curr. Opin. Neurobiol. 51, 60–69 (2018).
Google Scholar
Tedeschi, A. et al. ADF/cofilin-mediated actin turnover promotes axon regeneration in the adult CNS. Neuron 103, 1073–1085.e1076 (2019).
Google Scholar
Stern, S. et al. RhoA drives actin compaction to restrict axon regeneration and astrocyte reactivity after CNS injury. Neuron 109, 3436–3455.e9 (2021).
Google Scholar
Wu, D. et al. Chronic neuronal activation increases dynamic microtubules to enhance functional axon regeneration after dorsal root crush injury. Nat. Commun. 11, 6131 (2020).
Google Scholar
Hellal, F. et al. Microtubule stabilization reduces scarring and causes axon regeneration after spinal cord injury. Science 331, 928–931 (2011).
Google Scholar
Sengottuvel, V., Leibinger, M., Pfreimer, M., Andreadaki, A. & Fischer, D. Taxol facilitates axon regeneration in the mature CNS. J. Neurosci. 31, 2688–2699 (2011).
Google Scholar
Ruschel, J. et al. Systemic administration of epothilone B promotes axon regeneration after spinal cord injury. Science 348, 347–352 (2015).
Google Scholar
Brunden, K. R. et al. The characterization of microtubule-stabilizing drugs as possible therapeutic agents for Alzheimer’s disease and related tauopathies. Pharmacol. Res. 63, 341–351 (2011).
Google Scholar
Nettles, J. H. et al. The binding mode of epothilone A on alpha,beta-tubulin by electron crystallography. Science 305, 866–869 (2004).
Google Scholar
Prota, A. E. et al. Molecular mechanism of action of microtubule-stabilizing anticancer agents. Science 339, 587–590 (2013).
Google Scholar
Howes, S. C. et al. Structural differences between yeast and mammalian microtubules revealed by cryo-EM. J. Cell Biol. 216, 2669–2677 (2017).
Google Scholar
Perez, E. A. et al. Efficacy and safety of ixabepilone (BMS-247550) in a phase II study of patients with advanced breast cancer resistant to an anthracycline, a taxane, and capecitabine. J. Clin. Oncol. 25, 3407–3414 (2007).
Google Scholar
Colom, A. et al. A fluorescent membrane tension probe. Nat. Chem. 10, 1118–1125 (2018).
Google Scholar
Skaliora, I., Adams, R. & Blakemore, C. Morphology and growth patterns of developing thalamocortical axons. J. Neurosci. 20, 3650–3662 (2000).
Google Scholar
Goodson, H. V. & Jonasson, E. M. Microtubules and microtubule-associated proteins. Cold Spring Harb. Perspect. Biol. 10, a022608 (2018).
Google Scholar
Chretien, D., Metoz, F., Verde, F., Karsenti, E. & Wade, R. H. Lattice defects in microtubules: protofilament numbers vary within individual microtubules. J. Cell Biol. 117, 1031–1040 (1992).
Google Scholar
Mizuno, N. et al. Dynein and kinesin share an overlapping microtubule-binding site. EMBO J. 23, 2459–2467 (2004).
Google Scholar
Baas, P. W., Rao, A. N., Matamoros, A. J. & Leo, L. Stability properties of neuronal microtubules. Cytoskeleton 73, 442–460 (2016).
Google Scholar
Moores, C. A. et al. Mechanism of microtubule stabilization by doublecortin. Mol. Cell 14, 833–839 (2004).
Google Scholar
Tymanskyj, S. R. & Ma, L. MAP7 prevents axonal branch retraction by creating a stable microtubule boundary to rescue polymerization. J. Neurosci. 39, 7118–7131 (2019).
Google Scholar
Heidemann, S. R., Landers, J. M. & Hamborg, M. A. Polarity orientation of axonal microtubules. J. Cell Biol. 91, 661–665 (1981).
Google Scholar
Burton, P. R. & Paige, J. L. Polarity of axoplasmic microtubules in the olfactory nerve of the frog. Proc. Natl Acad. Sci. USA 78, 3269–3273 (1981).
Google Scholar
Baas, P. W. & Lin, S. Hooks and comets: the story of microtubule polarity orientation in the neuron. Dev. Neurobiol. 71, 403–418 (2011).
Google Scholar
Zhang, R., LaFrance, B. & Nogales, E. Separating the effects of nucleotide and EB binding on microtubule structure. Proc. Natl Acad. Sci. USA 115, E6191–E6200 (2018).
Google Scholar
Itzhak, D. N., Tyanova, S., Cox, J. & Borner, G. H. Global, quantitative and dynamic mapping of protein subcellular localization. eLife 5, e16950 (2016).
Google Scholar
Hiller, G. & Weber, K. Radioimmunoassay for tubulin: a quantitative comparison of the tubulin content of different established tissue culture cells and tissues. Cell 14, 795–804 (1978).
Google Scholar
Nedozralova, H. et al. In situ cryo-electron tomography reveals local cellular machineries for axon branch development. J. Cell Biol. 221, e202106086 (2022).
Google Scholar
Myers, K. A. & Baas, P. W. Kinesin-5 regulates the growth of the axon by acting as a brake on its microtubule array. J. Cell Biol. 178, 1081–1091 (2007).
Google Scholar
Cuveillier, C. et al. MAP6 is an intraluminal protein that induces neuronal microtubules to coil. Sci. Adv. 6, eaaz4344 (2020).
Google Scholar
Chakraborty, S. et al. Cryo-ET suggests tubulin chaperones form a subset of microtubule lumenal particles with a role in maintaining neuronal microtubules. Proc. Natl Acad. Sci. USA 122, e2404017121 (2025).
Google Scholar
Wang, Q., Crevenna, A. H., Kunze, I. & Mizuno, N. Structural basis for the extended CAP-Gly domains of p150(glued) binding to microtubules and the implication for tubulin dynamics. Proc. Natl Acad. Sci. USA 111, 11347–11352 (2014).
Google Scholar
Ayukawa, R. et al. GTP-dependent formation of straight tubulin oligomers leads to microtubule nucleation. J. Cell Biol. 220, e202007033 (2021).
Google Scholar
McIntosh, J. R. et al. Microtubules grow by the addition of bent guanosine triphosphate tubulin to the tips of curved protofilaments. J. Cell Biol. 217, 2691–2708 (2018).
Google Scholar
Mandelkow, E. M., Mandelkow, E. & Milligan, R. A. Microtubule dynamics and microtubule caps: a time-resolved cryo-electron microscopy study. J. Cell Biol. 114, 977–991 (1991).
Google Scholar
Ojeda-Lopez, M. A. et al. Transformation of Taxol-stabilized microtubules into inverted tubulin tubules triggered by a tubulin conformation switch. Nat. Mater. 13, 195–203 (2014).
Google Scholar
Basnet, N. et al. Direct induction of microtubule branching by microtubule nucleation factor SSNA1. Nat. Cell Biol. 20, 1172–1180 (2018).
Google Scholar
Zhang, B. et al. Synaptic vesicle size and number are regulated by a clathrin adaptor protein required for endocytosis. Neuron 21, 1465–1475 (1998).
Google Scholar
Silver, J., Schwab, M. E. & Popovich, P. G. Central nervous system regenerative failure: role of oligodendrocytes, astrocytes, and microglia. Cold Spring Harb. Perspect. Biol. 7, a020602 (2014).
Google Scholar
Richardson, P. M., McGuinness, U. M. & Aguayo, A. J. Axons from CNS neurons regenerate into PNS grafts. Nature 284, 264–265 (1980).
Google Scholar
Liu, K., Tedeschi, A., Park, K. K. & He, Z. Neuronal intrinsic mechanisms of axon regeneration. Annu. Rev. Neurosci. 34, 131–152 (2011).
Google Scholar
Cho, Y., Sloutsky, R., Naegle, K. M. & Cavalli, V. Injury-induced HDAC5 nuclear export is essential for axon regeneration. Cell 155, 894–908 (2013).
Google Scholar
Rishal, I. & Fainzilber, M. Axon-soma communication in neuronal injury. Nat. Rev. Neurosci. 15, 32–42 (2014).
Google Scholar
Varadarajan, S. G., Hunyara, J. L., Hamilton, N. R., Kolodkin, A. L. & Huberman, A. D. Central nervous system regeneration. Cell 185, 77–94 (2022).
Google Scholar
Tian, F. et al. Core transcription programs controlling injury-induced neurodegeneration of retinal ganglion cells. Neuron 110, 2607–2624.e2608 (2022).
Google Scholar
Moeendarbary, E. et al. The soft mechanical signature of glial scars in the central nervous system. Nat. Commun. 8, 14787 (2017).
Google Scholar
Giger, R. J., Hollis, E. R. 2nd & Tuszynski, M. H. Guidance molecules in axon regeneration. Cold Spring Harb. Perspect. Biol. 2, a001867 (2010).
Google Scholar
Gallo, V. & Deneen, B. Glial development: the crossroads of regeneration and repair in the CNS. Neuron 83, 283–308 (2014).
Google Scholar
Farias, G. G., Guardia, C. M., Britt, D. J., Guo, X. & Bonifacino, J. S. Sorting of dendritic and axonal vesicles at the pre-axonal exclusion zone. Cell Rep. 13, 1221–1232 (2015).
Google Scholar
Yang, R. et al. A novel strategy to visualize vesicle-bound kinesins reveals the diversity of kinesin-mediated transport. Traffic 20, 851–866 (2019).
Google Scholar
Dunn, S. et al. Differential trafficking of Kif5c on tyrosinated and detyrosinated microtubules in live cells. J. Cell Sci. 121, 1085–1095 (2008).
Google Scholar
Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
Google Scholar
Bodakuntla, S., Magiera, M. M. & Janke, C. Measuring the impact of tubulin posttranslational modifications on axonal transport. Methods Mol. Biol. 2101, 353–370 (2020).
Google Scholar
Hagen, W. J. H., Wan, W. & Briggs, J. A. G. Implementation of a cryo-electron tomography tilt-scheme optimized for high resolution subtomogram averaging. J. Struct. Biol. 197, 191–198 (2017).
Google Scholar
Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).
Google Scholar
Mastronarde, D. N. & Held, S. R. Automated tilt series alignment and tomographic reconstruction in IMOD. J. Struct. Biol. 197, 102–113 (2017).
Google Scholar
Tegunov, D. & Cramer, P. Real-time cryo-electron microscopy data preprocessing with Warp. Nat. Methods 16, 1146–1152 (2019).
Google Scholar
Scheres, S. H. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).
Google Scholar
Punjani, A., Rubinstein, J. L., Fleet, D. J. & Brubaker, M. A. cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat. Methods 14, 290–296 (2017).
Google Scholar
Grigorieff, N. Frealign: an exploratory tool for single-particle Cryo-EM. Methods Enzymol. 579, 191–226 (2016).
Google Scholar
Zhang, R. & Nogales, E. A new protocol to accurately determine microtubule lattice seam location. J. Struct. Biol. 192, 245–254 (2015).
Google Scholar
Meng, E. C. et al. UCSF ChimeraX: tools for structure building and analysis. Protein Sci. 32, e4792 (2023).
Google Scholar
Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D 66, 486–501 (2010).
Google Scholar
Afonine, P. V. et al. Real-space refinement in PHENIX for cryo-EM and crystallography. Acta Crystallogr. D 74, 531–544 (2018).
Google Scholar
Rusu, M., Starosolski, Z., Wahle, M., Rigort, A. & Wriggers, W. Automated tracing of filaments in 3D electron tomography reconstructions using Sculptor and Situs. J. Struct. Biol. 178, 121–128 (2012).
Google Scholar
Martinez-Sanchez, A. et al. Template-free detection and classification of membrane-bound complexes in cryo-electron tomograms. Nat. Methods 17, 209–216 (2020).
Google Scholar
Sousbie, T. The persistent cosmic web and its filamentary structure – I. Theory and implementation. Mon. Not. R. Astron. Soc. 414, 350–383 (2011).
Google Scholar
Comaniciu, D. & Meer, P. Mean shift: a robust approach toward feature space analysis. IEEE Trans. Pattern Anal. Mach. Intell. 24, 603–619 (2002).
Google Scholar
Digman, M. A., Caiolfa, V. R., Zamai, M. & Gratton, E. The phasor approach to fluorescence lifetime imaging analysis. Biophys. J. 94, L14–L16 (2008).
Google Scholar
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