The unique morphology and function of axons are sustained by the organization of the key elements of their cytoskeleton: microtubules, neurofilaments and actin.
Classical methods (electron microscopy and biochemistry) have been critical in identifying the morphology and composition of axonal cytostructures.
More recently, super-resolution microscopy, live-cell imaging and other new optical methods have been applied to the study of the axonal cytoskeleton.
This has led to major discoveries, in particular the existence of axonal actin structures such as rings, hot spots, trails and patches.
This Review summarizes the latest advances in our understanding of the axonal cytoskeleton and discusses key open questions in this field, such as the functions of newly discovered structures and the interplay between different cytoskeletal components.
The corporeal beauty of the neuronal cytoskeleton has captured the imagination of generations of scientists. One of the easiest cellular structures to visualize by light microscopy, its existence has been known for well over 100 years, yet we have only recently begun to fully appreciate its intricacy and diversity. Recent studies combining new probes with super-resolution microscopy and live imaging have revealed surprising details about the axonal cytoskeleton and, in particular, have discovered previously unknown actin-based structures. Along with traditional electron microscopy, these newer techniques offer a nanoscale view of the axonal cytoskeleton, which is important for our understanding of neuronal form and function, and lay the foundation for future studies. In this Review, we summarize existing concepts in the field and highlight contemporary discoveries that have fundamentally altered our perception of the axonal cytoskeleton.
This is a preview of subscription content, access via your institution
Open Access articles citing this article.
Proteomic and functional analyses of the periodic membrane skeleton in neurons
Nature Communications Open Access 09 June 2022
Fast widefield scan provides tunable and uniform illumination optimizing super-resolution microscopy on large fields
Nature Communications Open Access 24 May 2021
The model of local axon homeostasis - explaining the role and regulation of microtubule bundles in axon maintenance and pathology
Neural Development Open Access 09 November 2019
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Rent or buy this article
Get just this article for as long as you need it
Prices may be subject to local taxes which are calculated during checkout
Wilt, B. A. et al. Advances in light microscopy for neuroscience. Annu. Rev. Neurosci. 32, 435–506 (2009).
Maglione, M. & Sigrist, S. J. Seeing the forest tree by tree: super-resolution light microscopy meets the neurosciences. Nat. Neurosci. 16, 790–797 (2013).
Pannese, E. Neurocytology. (Springer, 2015).
Wuerker, R. B. & Kirkpatrick, J. B. Neuronal microtubules, neurofilaments, and microfilaments. Int. Rev. Cytol. 33, 45–75 (1972).
Frixione, E. The cytoskeleton of nerve cells in historic perspective. IBRO History of Neuroscience http://ibro.info/wp-content/uploads/2012/12/The-Cytoskeleton-of-Nerve-Cells-in-Historic-Perspective.pdf (2006).
Hirokawa, N. Cross-linker system between neurofilaments, microtubules, and membranous organelles in frog axons revealed by the quick-freeze, deep-etching method. J. Cell Biol. 94, 129–142 (1982).
Schnapp, B. J. & Reese, T. S. Cytoplasmic structure in rapid-frozen axons. J. Cell Biol. 94, 667–669 (1982). Using quick-freeze, deep-etching EM, references 6 and 7 provide the pictures that shaped our thinking about the axonal cytoskeleton for the following decades.
Shelanski, M. L. & Taylor, E. W. Isolation of a protein subunit from microtubules. J. Cell Biol. 34, 549–554 (1967).
Borisy, G. G. & Taylor, E. W. The mechanism of action of colchicine. Binding of colchincine-3H to cellular protein. J. Cell Biol. 34, 525–533 (1967).
Ishikawa, H., Bischoff, R. & Holtzer, H. Formation of arrowhead complexes with heavy meromyosin in a variety of cell types. J. Cell Biol. 43, 312–328 (1969).
Leterrier, C. The axon initial segment, 50 years later: a nexus for neuronal organization and function. Curr. Top. Membr. 77, 185–233 (2016).
Kohno, K. Neurotubules contained within the dendrite and axon of Purkinje cell of frog. Bull. Tokyo Dent. Univ. 11, 411–442 (1964).
Palay, S., Sotelo, C., Peters, A. & Orkand, P. The axon hillock and the initial segment. J. Cell Biol. 38, 193–201 (1968).
Peters, A., Proskauer, C. C. & Kaiserman-Abramof, I. R. The small pyramidal neuron of the rat cerebral cortex. The axon hillock and initial segment. J. Cell Biol. 39, 604–619 (1968).
Hirokawa, N. & Takemura, R. Molecular motors and mechanisms of directional transport in neurons. Nat. Rev. Neurosci. 6, 201–214 (2005).
Song, A.-H. et al. A selective filter for cytoplasmic transport at the axon initial segment. Cell 136, 1148–1160 (2009). This study identifies an actin-based sorting of vesicular trafficking at the axon entrance whose mechanism is still debated today.
Chan-Palay, V. The tripartite structure of the undercoat in initial segments of Purkinje cell axons. Z. Anat. Entwicklungsgesch 139, 1–10 (1972).
Rasband, M. N. The axon initial segment and the maintenance of neuronal polarity. Nat. Rev. Neurosci. 11, 552–562 (2010).
Kordeli, E., Lambert, S. & Bennett, V. AnkyrinG. A new ankyrin gene with neural-specific isoforms localized at the axonal initial segment and node of Ranvier. J. Biol. Chem. 270, 2352–2359 (1995).
Zhou, D. et al. AnkyrinG is required for clustering of voltage-gated Na channels at axon initial segments and for normal action potential firing. J. Cell Biol. 143, 1295–1304 (1998).
Berghs, S. et al. βIV spectrin, a new spectrin localized at axon initial segments and nodes of ranvier in the central and peripheral nervous system. J. Cell Biol. 151, 985–1002 (2000).
Komada, M. & Soriano, P. βIV-spectrin regulates sodium channel clustering through ankyrin-G at axon initial segments and nodes of Ranvier. J. Cell Biol. 156, 337–348 (2002).
Winckler, B., Forscher, P. & Mellman, I. A diffusion barrier maintains distribution of membrane proteins in polarized neurons. Nature 397, 698–701 (1999).
Nakada, C. et al. Accumulation of anchored proteins forms membrane diffusion barriers during neuronal polarization. Nat. Cell Biol. 5, 626–632 (2003). This article uses state-of-the-art single-particle tracking to demonstrate lipid immobilization at the AIS (in addition to protein immobilization shown in reference 23) and to probe the molecular basis of this diffusion barrier.
Hoffman, P. N. & Lasek, R. J. The slow component of axonal transport. Identification of major structural polypeptides of the axon and their generality among mammalian neurons. J. Cell Biol. 66, 351–366 (1975).
Black, M. M. & Lasek, R. J. Slow components of axonal transport: two cytoskeletal networks. J. Cell Biol. 86, 616–623 (1980).
Willard, M., Wiseman, M., Levine, J. & Skene, P. Axonal transport of actin in rabbit retinal ganglion cells. J. Cell Biol. 81, 581–591 (1979).
Galbraith, J. A. & Gallant, P. E. Axonal transport of tubulin and actin. J. Neurocytol. 29, 889–911 (2000).
Roy, S. Seeing the unseen: the hidden world of slow axonal transport. Neuroscientist 20, 71–81 (2014).
Wang, Y., Shyy, J. Y.-J. & Chien, S. Fluorescence proteins, live-cell imaging, and mechanobiology: seeing is believing. Annu. Rev. Biomed. Eng. 10, 1–38 (2008).
Kuczmarski, E. R. & Rosenbaum, J. L. Studies on the organization and localization of actin and myosin in neurons. J. Cell Biol. 80, 356–371 (1979).
Alonso, G., Gabrion, J., Travers, E. & Assenmacher, I. Ultrastructural organization of actin filaments in neurosecretory axons of the rat. Cell Tissue Res. 214, 323–341 (1981).
Nagele, R. G., Kosciuk, M. C., Hunter, E. T., Bush, K. T. & Lee, H. Immunoelectron microscopic localization of actin in neurites of cultured embryonic chick dorsal root ganglia: actin is a component of granular, microtubule-associated crossbridges. Brain Res. 474, 279–286 (1988).
Letourneau, P. C. Differences in the organization of actin in the growth cones compared with the neurites of cultured neurons from chick embryos. J. Cell Biol. 97, 963–973 (1983).
Letourneau, P. C. Actin in axons: stable scaffolds and dynamic filaments. Results Probl. Cell Differ. 48, 65–90 (2009).
Wang, L., Ho, C. L., Sun, D., Liem, R. K. & Brown, A. Rapid movement of axonal neurofilaments interrupted by prolonged pauses. Nat. Cell Biol. 2, 137–141 (2000).
Roy, S. et al. Neurofilaments are transported rapidly but intermittently in axons: implications for slow axonal transport. J. Neurosci. 20, 6849–6861 (2000). References 36 and 37 show rapid, intermittent movements of neurofilaments along axons, establishing the 'stop and go' model of neurofilament transport.
Stepanova, T. et al. Visualization of microtubule growth in cultured neurons via the use of EB3-GFP (end-binding protein 3-green fluorescent protein). J. Neurosci. 23, 2655–2664 (2003). This report describes pioneering the use of fluorescent EBs to detect the orientation of dynamic axonal microtubules, which is refined and extended in reference 39.
Yau, K. W. et al. Dendrites in vitro and in vivo contain microtubules of opposite polarity and axon formation correlates with uniform plus-end-out microtubule orientation. J. Neurosci. 36, 1071–1085 (2016).
Conde, C. & Caceres, A. Microtubule assembly, organization and dynamics in axons and dendrites. Nat. Rev. Neurosci. 10, 319–332 (2009).
Heidemann, S. R., Landers, J. M. & Hamborg, M. A. Polarity orientation of axonal microtubules. J. Cell Biol. 91, 661–665 (1981).
Baas, P. W., Deitch, J. S., Black, M. M. & Banker, G. A. Polarity orientation of microtubules in hippocampal neurons: uniformity in the axon and nonuniformity in the dendrite. Proc. Natl Acad. Sci. USA 85, 8335–8339 (1988).
Baas, P. W. & Lin, S. Hooks and comets: the story of microtubule polarity orientation in the neuron. Dev. Neurobiol. 71, 403–418 (2011).
van de Willige, D., Hoogenraad, C. C. & Akhmanova, A. Microtubule plus-end tracking proteins in neuronal development. Cell. Mol. Life Sci. 73, 2053–2077 (2016).
Kleele, T. et al. An assay to image neuronal microtubule dynamics in mice. Nat. Commun. 5, 4827 (2014).
Waxman, S. G. & Kocsis, J. D. The Axon. (Oxford Univ. Press, 1995).
Peters, A. & Vaughn, J. E. Microtubules and filaments in the axons and astrocytes of early postnatal rat optic nerves. J. Cell Biol. 32, 113–119 (1967).
Yu, W. & Baas, P. W. Changes in microtubule number and length during axon differentiation. J. Neurosci. 14, 2818–2829 (1994).
Bray, D. & Bunge, M. B. Serial analysis of microtubules in cultured rat sensory axons. J. Neurocytol. 10, 589–605 (1981).
Burton, P. R. Microtubules of frog olfactory axons: their length and number/axon. Brain Res. 409, 71–78 (1987).
Burton, P. R. Ultrastructural studies of microtubules and microtubule organizing centers of the vertebrate olfactory neuron. Microsc. Res. Tech. 23, 142–156 (1992).
Tsukita, S. & Ishikawa, H. The cytoskeleton in myelinated axons: serial section study. Biomed. Res. 2, 424–437 (1981).
Yogev, S., Cooper, R., Fetter, R., Horowitz, M. & Shen, K. Microtubule organization determines axonal transport dynamics. Neuron 92, 449–460 (2016). This article uses methods based on fluorescence and live-cell imaging to comprehensively map the organization of axonal microtubules in C. elegans neurons and correlate it to vesicular transport events.
Chalfie, M. & Thomson, J. N. Organization of neuronal microtubules in the nematode Caenorhabditis elegans. J. Cell Biol. 82, 278–289 (1979).
Mudrakola, H. V., Zhang, K. & Cui, B. Optically resolving individual microtubules in live axons. Structure 17, 1433–1441 (2009).
Mikhaylova, M. et al. Resolving bundled microtubules using anti-tubulin nanobodies. Nat. Commun. 6, 7933 (2015). This study demonstrates the need for small probes (nanobodies) in addition to super-resolution microscopy to probe the organization of microtubules within neuronal bundles.
van Coevorden-Hameete, M. H. et al. Antibodies to TRIM46 are associated with paraneoplastic neurological syndromes. Ann. Clin. Transl Neurol. 4, 680–686 (2017).
van Beuningen, S. F. B. et al. TRIM46 controls neuronal polarity and axon specification by driving the formation of parallel microtubule arrays. Neuron 88, 1208–1226 (2015).
Leterrier, C. & Dargent, B. No Pasaran! Role of the axon initial segment in the regulation of protein transport and the maintenance of axonal identity. Semin. Cell Dev. Biol. 27, 44–51 (2014).
Satake, T. et al. MTCL1 plays an essential role in maintaining Purkinje neuron axon initial segment. EMBO J. 36, 1227–1242 (2017).
Nakata, T. & Hirokawa, N. Microtubules provide directional cues for polarized axonal transport through interaction with kinesin motor head. J. Cell Biol. 162, 1045–1055 (2003).
Nakata, T., Niwa, S., Okada, Y., Perez, F. & Hirokawa, N. Preferential binding of a kinesin-1 motor to GTP-tubulin-rich microtubules underlies polarized vesicle transport. J. Cell Biol. 194, 245–255 (2011).
Leterrier, C. et al. End-binding proteins EB3 and EB1 link microtubules to ankyrin G in the axon initial segment. Proc. Natl Acad. Sci. USA 108, 8826–8831 (2011).
Freal, A. et al. Cooperative interactions between 480 kDa ankyrin-G and EB proteins assemble the axon initial segment. J. Neurosci. 36, 4421–4433 (2016).
Leterrier, C. et al. Nanoscale architecture of the axon initial segment reveals an organized and robust scaffold. Cell Rep. 13, 2781–2793 (2015). This article uses STORM to determine the architecture of the axon initial segment scaffold, highlighting the capability of super-resolution microscopy to delineate molecular complexes in neurons.
Yu, W., Centonze, V. E., Ahmad, F. J. & Baas, P. W. Microtubule nucleation and release from the neuronal centrosome. J. Cell Biol. 122, 349–359 (1993).
Ahmad, F. J., Joshi, H. C., Centonze, V. E. & Baas, P. W. Inhibition of microtubule nucleation at the neuronal centrosome compromises axon growth. Neuron 12, 271–280 (1994).
Baas, P. W. Microtubules and neuronal polarity: lessons from mitosis. Neuron 22, 23–31 (1999).
Stiess, M. et al. Axon extension occurs independently of centrosomal microtubule nucleation. Science 327, 704–707 (2010).
Nguyen, M. M., Stone, M. C. & Rolls, M. M. Microtubules are organized independently of the centrosome in Drosophila neurons. Neural Dev. 6, 38 (2011).
Yonezawa, S., Shigematsu, M., Hirata, K. & Hayashi, K. Loss of γ-tubulin, GCP-WD/NEDD1 and CDK5RAP2 from the centrosome of neurons in developing mouse cerebral and cerebellar cortex. Acta Histochem. Cytochem. 48, 145–152 (2015).
Yau, K. W. et al. Microtubule minus-end binding protein CAMSAP2 controls axon specification and dendrite development. Neuron 82, 1058–1073 (2014).
Kuijpers, M. & Hoogenraad, C. C. Centrosomes, microtubules and neuronal development. Mol. Cell. Neurosci. 48, 349–358 (2011).
Ori-McKenney, K. M., Jan, L. Y. & Jan, Y.-N. Golgi outposts shape dendrite morphology by functioning as sites of acentrosomal microtubule nucleation in neurons. Neuron 76, 921–930 (2012).
Delandre, C., Amikura, R. & Moore, A. W. Microtubule nucleation and organization in dendrites. Cell Cycle 15, 1685–1692 (2016).
Sánchez-Huertas, C. et al. Non-centrosomal nucleation mediated by augmin organizes microtubules in post-mitotic neurons and controls axonal microtubule polarity. Nat. Commun. 7, 12187 (2016).
Kapitein, L. C. & Hoogenraad, C. C. Building the neuronal microtubule cytoskeleton. Neuron 87, 492–506 (2015).
Howard, J. & Hyman, A. A. Growth, fluctuation and switching at microtubule plus ends. Nat. Rev. Mol. Cell Biol. 10, 569–574 (2009).
Akhmanova, A. & Hoogenraad, C. C. Microtubule minus-end-targeting proteins. Curr. Biol. 25, R162–R171 (2015).
Wu, J. & Akhmanova, A. Microtubule-organizing centers. Annu. Rev. Cell Dev. Biol. http://dx.doi.org/10.1146/annurev-cellbio-100616-060615 (2017).
Goodwin, S. S. & Vale, R. D. Patronin regulates the microtubule network by protecting microtubule minus ends. Cell 143, 263–274 (2010).
Richardson, C. E. et al. PTRN-1, a microtubule minus end-binding CAMSAP homolog, promotes microtubule function in Caenorhabditis elegans neurons. eLife 3, e01498 (2014).
Marcette, J. D., Chen, J. J. & Nonet, M. L. The Caenorhabditis elegans microtubule minus-end binding homolog PTRN-1 stabilizes synapses and neurites. eLife 3, e01637 (2014).
Baas, P. W., Rao, A. N., Matamoros, A. J. & Leo, L. Stability properties of neuronal microtubules. Cytoskeleton 73, 442–460 (2016).
Baas, P. W., Slaughter, T., Brown, A. & Black, M. M. Microtubule dynamics in axons and dendrites. J. Neurosci. Res. 30, 134–153 (1991).
Baas, P. W. & Black, M. M. Individual microtubules in the axon consist of domains that differ in both composition and stability. J. Cell Biol. 111, 495–509 (1990).
Ahmad, F. J., Pienkowski, T. P. & Baas, P. W. Regional differences in microtubule dynamics in the axon. J. Neurosci. 13, 856–866 (1993).
Janke, C. & Bulinski, J. C. Post-translational regulation of the microtubule cytoskeleton: mechanisms and functions. Nat. Rev. Mol. Cell Biol. 12, 773–786 (2011).
Song, Y. et al. Transglutaminase and polyamination of tubulin: posttranslational modification for stabilizing axonal microtubules. Neuron 78, 109–123 (2013).
Sirajuddin, M., Rice, L. M. & Vale, R. D. Regulation of microtubule motors by tubulin isotypes and post-translational modifications. Nat. Cell Biol. 16, 335–344 (2014).
Janke, C. & Kneussel, M. Tubulin post-translational modifications: encoding functions on the neuronal microtubule cytoskeleton. Trends Neurosci. 33, 362–372 (2010).
Xu, Z. et al. Microtubules acquire resistance from mechanical breakage through intralumenal acetylation. Science 356, 328–332 (2017).
Hammond, J. W. et al. Posttranslational modifications of tubulin and the polarized transport of kinesin-1 in neurons. Mol. Biol. Cell 21, 572–583 (2010).
Konishi, Y. & Setou, M. Tubulin tyrosination navigates the kinesin-1 motor domain to axons. Nat. Neurosci. 12, 559–567 (2009).
Tapia, M., Wandosell, F. & Garrido, J. J. Impaired function of HDAC6 slows down axonal growth and interferes with axon initial segment development. PLoS ONE 5, e12908 (2010).
Dimitrov, A. et al. Detection of GTP-tubulin conformation in vivo reveals a role for GTP remnants in microtubule rescues. Science 322, 1353–1356 (2008).
Cassimeris, L., Guglielmi, L., Denis, V., Larroque, C. & Martineau, P. Specific in vivo labeling of tyrosinated α-tubulin and measurement of microtubule dynamics using a GFP tagged, cytoplasmically expressed recombinant antibody. PLoS ONE 8, e59812 (2013).
Borowiak, M. et al. Photoswitchable inhibitors of microtubule dynamics optically control mitosis and cell death. Cell 162, 403–411 (2015).
Tashiro, T. & Komiya, Y. Organization and slow axonal transport of cytoskeletal proteins under normal and regenerating conditions. Mol. Neurobiol. 6, 301–311 (1992).
Hoffman, P. N., Lopata, M. A., Watson, D. F. & Luduena, R. F. Axonal transport of class II and III β-tubulin: evidence that the slow component wave represents the movement of only a small fraction of the tubulin in mature motor axons. J. Cell Biol. 119, 595–604 (1992).
Terasaki, M., Schmidek, A., Galbraith, J. A., Gallant, P. E. & Reese, T. S. Transport of cytoskeletal elements in the squid giant axon. Proc. Natl Acad. Sci. USA 92, 11500–11503 (1995).
Dent, E. W., Callaway, J. L., Szebenyi, G., Baas, P. W. & Kalil, K. Reorganization and movement of microtubules in axonal growth cones and developing interstitial branches. J. Neurosci. 19, 8894–8908 (1999).
Galbraith, J. A., Reese, T. S., Schlief, M. L. & Gallant, P. E. Slow transport of unpolymerized tubulin and polymerized neurofilament in the squid giant axon. Proc. Natl Acad. Sci. USA 96, 11589–11594 (1999).
Terada, S., Kinjo, M. & Hirokawa, N. Oligomeric tubulin in large transporting complex is transported via kinesin in squid giant axons. Cell 103, 141–155 (2000).
Wang, L. & Brown, A. Rapid movement of microtubules in axons. Curr. Biol. 12, 1496–1501 (2002).
Brown, A. Axonal transport of membranous and nonmembranous cargoes: a unified perspective. J. Cell Biol. 160, 817–821 (2003).
He, Y. et al. Role of cytoplasmic dynein in the axonal transport of microtubules and neurofilaments. J. Cell Biol. 168, 697–703 (2005).
Rao, A. N. et al. Cytoplasmic dynein transports axonal microtubules in a polarity-sorting manner. Cell Rep. 19, 2210–2219 (2017).
Ganguly, A. et al. Hsc70 chaperone activity is required for the cytosolic slow axonal transport of synapsin. J. Cell Biol. 216, 2059–2074 (2017).
Jolly, A. L. et al. Kinesin-1 heavy chain mediates microtubule sliding to drive changes in cell shape. Proc. Natl Acad. Sci. USA 107, 12151–12156 (2010).
Keating, T. J., Peloquin, J. G., Rodionov, V. I., Momcilovic, D. & Borisy, G. G. Microtubule release from the centrosome. Proc. Natl Acad. Sci. USA 94, 5078–5083 (1997).
Hirokawa, N., Glicksman, M. A. & Willard, M. B. Organization of mammalian neurofilament polypeptides within the neuronal cytoskeleton. J. Cell Biol. 98, 1523–1536 (1984).
Laser-Azogui, A., Kornreich, M., Malka-Gibor, E. & Beck, R. Neurofilament assembly and function during neuronal development. Curr. Opin. Cell Biol. 32, 92–101 (2015).
Uchida, A., Colakoglu, G., Wang, L., Monsma, P. C. & Brown, A. Severing and end-to-end annealing of neurofilaments in neurons. Proc. Natl Acad. Sci. USA 110, E2696–E2705 (2013).
Leduc, C. & Etienne-Manneville, S. Regulation of microtubule-associated motors drives intermediate filament network polarization. J. Cell Biol. 216, 1689–1703 (2017).
Black, M. M. & Lasek, R. J. Axonal transport of actin: slow component b is the principal source of actin for the axon. Brain Res. 171, 401–413 (1979).
McQuarrie, I. G., Brady, S. T. & Lasek, R. J. Diversity in the axonal transport of structural proteins: major differences between optic and spinal axons in the rat. J. Neurosci. 6, 1593–1605 (1986).
Nixon, R. A. & Logvinenko, K. B. Multiple fates of newly synthesized neurofilament proteins: evidence for a stationary neurofilament network distributed nonuniformly along axons of retinal ganglion cell neurons. J. Cell Biol. 102, 647–659 (1986).
Hirokawa, N., Funakoshi, S. T. & Takeda, S. Slow axonal transport: the subunit transport model. Trends Cell Biol. 7, 384–388 (1997).
Baas, P. W. & Brown, A. Slow axonal transport: the polymer transport model. Trends Cell Biol. 7, 380–384 (1997). References 119 and 120 offer a summary of the 1990s debate on the mechanism of slow axonal transport for cytoskeletal components.
Yan, Y. & Brown, A. Neurofilament polymer transport in axons. J. Neurosci. 25, 7014–7021 (2005).
Uchida, A., Alami, N. H. & Brown, A. Tight functional coupling of kinesin-1A and dynein motors in the bidirectional transport of neurofilaments. Mol. Biol. Cell 20, 4997–5006 (2009).
LeBeux, Y. J. & Willemot, J. An ultrastructural study of the microfilaments in rat brain by means of heavy meromyosin labeling. I. The perikaryon, the dendrites and the axon. Cell Tissue Res. 160, 1–36 (1975).
Fath, K. R. & Lasek, R. J. Two classes of actin microfilaments are associated with the inner cytoskeleton of axons. J. Cell Biol. 107, 613–621 (1988).
Bearer, E. L. & Reese, T. S. Association of actin filaments with axonal microtubule tracts. J. Neurocytol. 28, 85–98 (1999).
Spooner, B. S. & Holladay, C. R. Distribution of tubulin and actin in neurites and growth cones of differentiating nerve cells. Cytoskeleton 1, 167–178 (1981).
Xu, K., Zhong, G. & Zhuang, X. Actin, spectrin, and associated proteins form a periodic cytoskeletal structure in axons. Science 339, 452–456 (2013). This study identifies the periodic organization of submembrane axonal actin, with actin rings regularly spaced every 190 nm by spectrins.
Bennett, V., Davis, J. Q. & Fowler, W. E. Brain spectrin, a membrane-associated protein related in structure and function to erythrocyte spectrin. Nature 299, 126–131 (1982).
Glenney, J. R., Glenney, P. & Weber, K. F-Actin-binding and cross-linking properties of porcine brain fodrin, a spectrin-related molecule. J. Biol. Chem. 257, 9781–9787 (1982).
Lukinavicius, G. et al. Fluorogenic probes for live-cell imaging of the cytoskeleton. Nat. Methods 11, 731–733 (2014).
D'Este, E., Kamin, D., Göttfert, F., El-Hady, A. & Hell, S. W. STED nanoscopy reveals the ubiquity of subcortical cytoskeleton periodicity in living neurons. Cell Rep. 10, 1246–1251 (2015).
D'Este, E., Kamin, D., Balzarotti, F. & Hell, S. W. Ultrastructural anatomy of nodes of Ranvier in the peripheral nervous system as revealed by STED microscopy. Proc. Natl Acad. Sci. USA 114, E191–E199 (2016).
D'Este, E. et al. Subcortical cytoskeleton periodicity throughout the nervous system. Sci. Rep. 6, 22741 (2016).
He, J. et al. Prevalent presence of periodic actin-spectrin-based membrane skeleton in a broad range of neuronal cell types and animal species. Proc. Natl Acad. Sci. USA 113, 6029–6034 (2016).
Qu, Y., Hahn, I., Webb, S. E. D. & Prokop, A. Periodic actin structures in neuronal axons are required to maintain microtubules. Mol. Biol. Cell 28, 296–308 (2016).
Sidenstein, S. C. et al. Multicolour multilevel STED nanoscopy of actin/spectrin organization at synapses. Sci. Rep. 6, 26725 (2016).
Bär, J., Kobler, O., van Bommel, B. & Mikhaylova, M. Periodic F-actin structures shape the neck of dendritic spines. Sci. Rep. 6, 37136 (2016).
Han, B., Zhou, R., Xia, C. & Zhuang, X. Structural organization of the actin-spectrin-based membrane skeleton in dendrites and soma of neurons. Proc. Natl Acad. Sci. USA 114, E6678–E6685 (2017).
Zhong, G. et al. Developmental mechanism of the periodic membrane skeleton in axons. eLife 3, 194 (2014).
Leite, S. C. et al. The actin-binding protein α-adducin is required for maintaining axon diameter. Cell Rep. 15, 490–498 (2016). References 135, 139 and 140 present the first information about the mechanisms of actin ring assembly and maintenance.
Leite, S. C. & Sousa, M. M. The neuronal and actin commitment: why do neurons need rings? Cytoskeleton 73, 424–434 (2016).
Galiano, M. R. et al. A distal axonal cytoskeleton forms an intra-axonal boundary that controls axon initial segment assembly. Cell 149, 1125–1139 (2012).
Watanabe, K. et al. Networks of polarized actin filaments in the axon initial segment provide a mechanism for sorting axonal and dendritic proteins. Cell Rep. 2, 1546–1553 (2012). This study identifies actin patches inside the AIS and, together with reference 245, proposed that they regulate vesicular entrance into the axon.
Jones, S. L., Korobova, F. & Svitkina, T. Axon initial segment cytoskeleton comprises a multiprotein submembranous coat containing sparse actin filaments. J. Cell Biol. 205, 67–81 (2014).
Al-Bassam, S., Xu, M., Wandless, T. J. & Arnold, D. B. Differential trafficking of transport vesicles contributes to the localization of dendritic proteins. Cell Rep. 2, 89–100 (2012).
Janssen, A. F. J. et al. Myosin-V induces cargo immobilization and clustering at the axon initial segment. Front. Cell Neurosci. 11, 89 (2017).
Nirschl, J. J., Ghiretti, A. E. & Holzbaur, E. L. F. The impact of cytoskeletal organization on the local regulation of neuronal transport. Nat. Rev. Neurosci. 18, 585–597 (2017). This recent review gives a contemporary view of local regulation of the transport machinery in axons, dendrites and synapses.
Ganguly, A. et al. A dynamic formin-dependent deep F-actin network in axons. J. Cell Biol. 104, 20576–20417 (2015). This study presents the discovery of new actin structures within axons: static, intermittent hot spots and dynamic filamentous trails spurting from them.
Ruthel, G. & Banker, G. A. Actin-dependent anterograde movement of growth-cone-like structures along growing hippocampal axons: a novel form of axonal transport? Cell. Motil. Cytoskeleton 40, 160–173 (1998).
Flynn, K. C. et al. ADF/Cofilin-mediated actin retrograde flow directs neurite formation in the developing brain. Neuron 76, 1091–1107 (2012).
Tint, I., Jean, D., Baas, P. W. & Black, M. M. Doublecortin associates with microtubules preferentially in regions of the axon displaying actin-rich protrusive structures. J. Neurosci. 29, 10995–11010 (2009).
Katsuno, H. & Sakumura, Y. Actin migration driven by directional assembly and disassembly of membrane-anchored actin filaments. Cell Rep. 12, 648–660 (2015). This article combines live-cell imaging, micromanipulation and force measurements to detail the organization and mechanisms of axonal actin waves.
Winans, A. M., Collins, S. R. & Meyer, T. Waves of actin and microtubule polymerization drive microtubule-based transport and neurite growth before single axon formation. eLife 5, e12387 (2016).
Tomba, C. et al. Geometrical determinants of neuronal actin waves. Front. Cell Neurosci. 11, 86 (2017).
Flynn, K. C., Pak, C. W., Shaw, A. E., Bradke, F. & Bamburg, J. R. Growth cone-like waves transport actin and promote axonogenesis and neurite branching. Dev. Neurobiol. 69, 761–779 (2009).
Roy, S. Waves, rings, and trails: the scenic landscape of axonal actin. J. Cell Biol. 212, 131–134 (2016).
Allard, J. & Mogilner, A. Traveling waves in actin dynamics and cell motility. Curr. Opin. Cell Biol. 25, 107–115 (2013).
Arnold, D. B. & Gallo, G. Structure meets function: actin filaments and myosin motors in the axon. J. Neurochem. 129, 213–220 (2013).
Spillane, M. et al. The actin nucleating Arp2/3 complex contributes to the formation of axonal filopodia and branches through the regulation of actin patch precursors to filopodia. Dev. Neurobiol. 71, 747–758 (2011).
Armijo-Weingart, L. & Gallo, G. It takes a village to raise a branch: Cellular mechanisms of the initiation of axon collateral branches. Mol. Cell. Neurosci. http://dx.doi.org/10.1016/j.mcn.2017.03.007 (2017).
Chetta, J., Love, J. M., Bober, B. G. & Shah, S. B. Bidirectional actin transport is influenced by microtubule and actin stability. Cell. Mol. Life Sci. 72, 4205–4220 (2015).
Rogers, S. L. & Gelfand, V. I. Myosin cooperates with microtubule motors during organelle transport in melanophores. Curr. Biol. 8, 161–164 (1998).
Bridgman, P. C. Myosin-dependent transport in neurons. J. Neurobiol. 58, 164–174 (2004).
Lai, L. & Cao, J. Spectrins in axonal cytoskeletons: dynamics revealed by extensions and fluctuations. J. Chem. Phys. 141, 015101 (2014).
Zhang, Y. et al. Modeling of the axon membrane skeleton structure and implications for its mechanical properties. PLoS Comput. Biol. 13, e1005407 (2017).
Hammarlund, M., Jorgensen, E. M. & Bastiani, M. J. Axons break in animals lacking beta-spectrin. J. Cell Biol. 176, 269–275 (2007).
Krieg, M. et al. Genetic defects in β-spectrin and tau sensitize C. elegans axons to movement-induced damage via torque-tension coupling. eLife 6, 1187 (2017).
Krieg, M., Dunn, A. R. & Goodman, M. B. Mechanical control of the sense of touch by β-spectrin. Nat. Cell Biol. 16, 224–233 (2014). References 167 and 168 combine in vivo force measurements, mutant analysis, super-resolution microscopy and modelling to explore the role of microtubules and the actin–spectrin submembrane scaffold in the mechanical robustness of axons.
Stephan, R. et al. Hierarchical microtubule organization controls axon caliber and transport and determines synaptic structure and stability. Dev. Cell 33, 5–21 (2015).
Taylor, A. M., Dieterich, D. C., Ito, H. T., Kim, S. A. & Schuman, E. M. Microfluidic local perfusion chambers for the visualization and manipulation of synapses. Neuron 66, 57–68 (2010).
Harterink, M. et al. DeActs: genetically encoded tools for perturbing the actin cytoskeleton in single cells. Nat. Methods 14, 479–482 (2017).
Albrecht, D. et al. Nanoscopic compartmentalization of membrane protein motion at the axon initial segment. J. Cell Biol. 215, 37–46 (2016). This study combines single-particle tracking and super-resolution microscopy to detail how the AIS surface diffusion barrier forms and operates.
Brachet, A. et al. Ankyrin G restricts ion channel diffusion at the axonal initial segment before the establishment of the diffusion barrier. J. Cell Biol. 191, 383–395 (2010).
Dance, A. Inner workings: uncovering the neuron's internal skeleton. Proc. Natl Acad. Sci. USA 113, 13931–13933 (2016).
Tilney, L. G. & Portnoy, D. A. Actin filaments and the growth, movement, and spread of the intracellular bacterial parasite, Listeria monocytogenes. J. Cell Biol. 109, 1597–1608 (1989).
Wagner, O. I. et al. Mechanisms of mitochondria-neurofilament interactions. J. Neurosci. 23, 9046–9058 (2003).
Schermelleh, L., Heintzmann, R. & Leonhardt, H. A guide to super-resolution fluorescence microscopy. J. Cell Biol. 190, 165–175 (2010).
Toomre, D. & Bewersdorf, J. A new wave of cellular imaging. Annu. Rev. Cell Dev. Biol. 26, 285–314 (2010).
Fornasiero, E. F. & Opazo, F. Super-resolution imaging for cell biologists. Bioessays 37, 436–451 (2015).
Rust, M. J., Bates, M. & Zhuang, X. Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat. Methods 3, 793–795 (2006).
Betzig, E. et al. Imaging intracellular fluorescent proteins at nanometer resolution. Science 313, 1642–1645 (2006).
Gustafsson, M. G. Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy. J. Microsc. 198, 82–87 (2000).
Gustafsson, M. G. L. Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution. Proc. Natl Acad. Sci. USA 102, 13081–13086 (2005).
Klar, T. A., Jakobs, S., Dyba, M., Egner, A. & Hell, S. W. Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission. Proc. Natl Acad. Sci. USA 97, 8206–8210 (2000).
Dotti, C. G., Sullivan, C. & Banker, G. A. The establishment of polarity by hippocampal neurons in culture. J. Neurosci. 8, 1454–1468 (1988).
Dent, E. W. et al. Filopodia are required for cortical neurite initiation. Nat. Cell Biol. 9, 1347–1359 (2007).
Flynn, K. C. The cytoskeleton and neurite initiation. BioArchitecture 3, 86–109 (2014).
Jacobson, C., Schnapp, B. & Banker, G. A. A. Change in the selective translocation of the kinesin-1 motor domain marks the initial specification of the axon. Neuron 49, 797–804 (2006).
Randlett, O., Poggi, L., Zolessi, F. R. & Harris, W. A. The oriented emergence of axons from retinal ganglion cells is directed by laminin contact in vivo. Neuron 70, 266–280 (2011).
Lewis, T. L., Courchet, J. & Polleux, F. Cell biology in neuroscience: cellular and molecular mechanisms underlying axon formation, growth, and branching. J. Cell Biol. 202, 837–848 (2013).
Bentley, M. & Banker, G. A. The cellular mechanisms that maintain neuronal polarity. Nat. Rev. Neurosci. 17, 611–622 (2016).
Schelski, M. & Bradke, F. Neuronal polarization: from spatiotemporal signaling to cytoskeletal dynamics. Mol. Cell. Neurosci. http://dx.doi.org/10.1016/j.mcn.2017.03.008. (2017).
Bradke, F. & Dotti, C. G. Neuronal polarity: vectorial cytoplasmic flow precedes axon formation. Neuron 19, 1175–1186 (1997).
Bradke, F. & Dotti, C. G. The role of local actin instability in axon formation. Science 283, 1931–1934 (1999).
Witte, H., Neukirchen, D. & Bradke, F. Microtubule stabilization specifies initial neuronal polarization. J. Cell Biol. 180, 619–632 (2008).
Omotade, O. F., Pollitt, S. L. & Zheng, J. Q. Actin-based growth cone motility and guidance. Mol. Cell. Neurosci. http://dx.doi.org/10.1016/j.mcn.2017.03.001 (2017).
Vitriol, E. A. & Zheng, J. Q. Growth cone travel in space and time: the cellular ensemble of cytoskeleton, adhesion, and membrane. Neuron 73, 1068–1081 (2012).
Dent, E. W., Gupton, S. L. & Gertler, F. B. The growth cone cytoskeleton in axon outgrowth and guidance. Cold Spring Harb. Perspect. Biol. 3, a001800 (2011).
Nozumi, M., Nakatsu, F., Katoh, K. & Igarashi, M. Coordinated movement of vesicles and actin bundles during nerve growth revealed by superresolution microscopy. Cell Rep. 18, 2203–2216 (2017).
Hedstrom, K. et al. Neurofascin assembles a specialized extracellular matrix at the axon initial segment. J. Cell Biol. 178, 875–886 (2007).
Boiko, T. et al. Ankyrin-dependent and -independent mechanisms orchestrate axonal compartmentalization of L1 family members neurofascin and L1/neuron-glia cell adhesion molecule. J. Neurosci. 27, 590–603 (2007).
Kalil, K. & Dent, E. W. Branch management: mechanisms of axon branching in the developing vertebrate CNS. Nat. Rev. Neurosci. 15, 7–18 (2014).
McAllister, A. Dynamic aspects of CNS synapse formation. Annu. Rev. Neurosci. 30, 425–450 (2007).
Work in the Leterrier laboratory is supported by the Centre National de la Recherche Scientifique (CNRS) Action Thématique et Incitative sur Programme (ATIP)–Avenir programme AO2016. Work in the Roy laboratory is supported by US National Institutes of Health (NIH) grants R01NS075233, R01AG048218 and R21 AG052404.
The authors declare no competing financial interests.
- Electron microscopy
A group of methods that generate an image of a sample by using a beam of electrons. Electrons can be detected after passing through the sample (transmission electron microscopy) or after being reflected (scanning electron microscopy). Electron microscopy can routinely reach 1 nm resolution (the size of a single amino acid) but usually requires the sample to be placed in a vacuum and is therefore destructive and most applicable to fixed samples labelled with specific procedures.
- Immuno-electron microscopy
An electron microscopy modality in which proteins labelled by using antibodies that are tagged with small gold beads are imaged, allowing their localization.
- Microtubule plus-end-tracking proteins
A set of proteins that bind the growing plus ends of microtubules. The core components of this complex are end-binding proteins, dimeric proteins that interact with the specific tubulin conformation found at the plus end.
- Moiré effect
The emergence of a third pattern due to the superposition of two patterns with distinct frequencies. In microscopy, this effect is exploited in structured illumination microscopy by illuminating the sample with periodic patterns of light and using the resulting Moiré pattern to infer sample details that are beyond the diffraction limit.
An organelle that nucleates and controls the organization of microtubules and regulates cell-cycle progression.
- Microtubule-associated proteins
(MAPs). The repertoire of proteins that bind to microtubules. They can associate with the microtubule lattice or with the minus end or plus end of the microtubules. Microtubule-associated molecular motor complexes are also MAPs.
- Fluorescence recovery after photobleaching
A method used to measure the diffusion or transport of molecules. It requires tagging of the molecule of interest with a fluorescent marker, photobleaching of the label with a pulse of laser light and a subsequent measure of the rate of fluorescence recovery into the bleached area as other labelled molecules move into it.
Of a method to manipulate cells (for seeding, incubation or labelling) at the sub-millimetre scale by using small volumes of medium that are pumped into miniaturized culturing devices.
- En-passant boutons
Presynaptic specializations along axons that make contact with downstream neurons, as opposed to synaptic terminals at the extremity of axons. In hippocampal and cortical neuronal cultures, most presynapses are en-passant boutons.
Rights and permissions
About this article
Cite this article
Leterrier, C., Dubey, P. & Roy, S. The nano-architecture of the axonal cytoskeleton. Nat Rev Neurosci 18, 713–726 (2017). https://doi.org/10.1038/nrn.2017.129
This article is cited by
Proteomic and functional analyses of the periodic membrane skeleton in neurons
Nature Communications (2022)
Advanced imaging and labelling methods to decipher brain cell organization and function
Nature Reviews Neuroscience (2021)
Fast widefield scan provides tunable and uniform illumination optimizing super-resolution microscopy on large fields
Nature Communications (2021)
Deuterium double quantum-filtered NMR studies of peripheral and optic nerves
Magnetic Resonance Materials in Physics, Biology and Medicine (2021)
Imaging of spine synapses using super-resolution microscopy
Anatomical Science International (2021)