CoChat
Sign Up
Cited by 45Cellular Mechanics and InteractionsAxon Guidance and Neuronal SignalingProtein Kinase Regulation and GTPase Signaling

The DOI matches its Crossref record; 2 of the 2 references it cites were found in a registry or an index.

Cofilactin filaments regulate filopodial structure and dynamics in neuronal growth cones

Open access
Hylton, Ryan K., Heebner, Jessica E., Grillo, Michael A., Swulius, Matthew T.·2022·Nature Communications
DOI 10.1038/s41467-022-30116-xPMC PMC9068697
Open this paper in CoChat — free accountOpen PDF

Abstract

Cofilin is best known for its ability to sever actin filaments and facilitate cytoskeletal recycling inside of cells, but at higher concentrations in vitro, cofilin stabilizes a more flexible, hyper-twisted state of actin known as “cofilactin”. While this filament state is well studied, a structural role for cofilactin in dynamic cellular processes has not been observed. With a combination of cryo-electron tomography and fluorescence imaging in neuronal growth cones, we observe that filopodial actin filaments switch between a fascin-linked and a cofilin-decorated state, and that cofilactin is associated with a variety of dynamic events within filopodia. The switch to cofilactin filaments occurs in a graded fashion and correlates with a decline in fascin cross-linking within the filopodia, which is associated with curvature in the bundle. Our tomographic data reveal that the hyper-twisting of actin from cofilin binding leads to a rearrangement of filament packing, which largely excludes fascin from the base of filopodia. Our results provide mechanistic insight into the fundamentals of cytoskeletal remodeling inside of confined cellular spaces, and how the interplay between fascin and cofilin regulates the dynamics of searching filopodia.

Overview

The claims and statistics of this paper haven’t been checked yet.

Run the check — free, no account needed. It reads the open-access text and checks the sources it cites.

A few free checks a day · papers already verified open instantly · results are public

Integrity signals

Registry

Crossref · 10.1038/s41467-022-30116-x

Cites

2 of 2 references verified0 citation errors · 0 not checkable

Statistics

Verification

Claims the paper makes

The claims and statistics of this paper haven’t been checked yet.

Run the check — free, no account needed. It reads the open-access text and checks the sources it cites.

A few free checks a day · papers already verified open instantly · results are public

Claims other papers make about this one

Each is checked against this paper’s own text: does it say what the citing paper says it does?

Self-check

Percentages, sums and reported differences re-computed from the paper’s own numbers.

The self-check needs the paper’s full text.

Citations

Cites

Flagged first, then by citations

Show all 2 references

Research works printed but not linked: 70

  1. Kerstein, P. C., Nichol, R. H. & Gomez, T. M. Mechanochemical regulation of growth cone motility. Front. Cell Neurosci. 9, 244 (2015). · DOI
  2. Lowery, L. A. & Vactor, D. V. The trip of the tip: understanding the growth cone machinery. Nat. Rev. Mol. Cell Biol. 10, 332–343 (2009). · DOI
  3. 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). · DOI
  4. Stoeckli, E. T. Understanding axon guidance: are we nearly there yet? Development 145, (2018). · DOI · Paper page
  5. 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). · DOI
  6. Flynn, K. C. et al. ADF/cofilin-mediated actin retrograde flow directs neurite formation in the developing brain. Neuron 76, 1091–1107 (2012). · DOI
  7. Gomez, T. M. & Letourneau, P. C. Actin dynamics in growth cone motility and navigation. J. Neurochem. 129, 221–234 (2014). · DOI
  8. Omotade, O. F., Pollitt, S. L. & Zheng, J. Q. Actin-based growth cone motility and guidance. Mol. Cell. Neurosci. 84, 4–10 (2017). · DOI
  9. Tamariz, E. & Varela-Echavarría, A. The discovery of the growth cone and its influence on the study of axon guidance. Front. Neuroanat. 9, 51 (2015). · DOI
  10. Gasperini, R. J. et al. How does calcium interact with the cytoskeleton to regulate growth cone motility during axon pathfinding? Mol. Cell. Neurosci. 84, 29–35 (2017). · DOI
  11. Hall, A. & Lalli, G. Rho and Ras GTPases in axon growth, guidance, and branching. Cold Spring Harb. Perspect. Biol. 2, a001818 (2010). · DOI · Paper page
  12. Henley, J. & Poo, M. Guiding neuronal growth cones using Ca2+ signals. Trends Cell Biol. 14, 320–330 (2004). · DOI
  13. Piper, M., Horck, Fvan & Holt, C. The role of cyclic nucleotides in axon guidance. Adv. Exp. Med. Biol. 621, 134–143 (2007). · DOI
  14. Tojima, T., Hines, J. H., Henley, J. R. & Kamiguchi, H. Second messengers and membrane trafficking direct and organize growth cone steering. Nat. Rev. Neurosci. 12, 191–203 (2011). · DOI
  15. Wen, Z., Guirland, C., Ming, G. & Zheng, J. Q. A CaMKII/Calcineurin switch controls the direction of Ca2+-dependent growth cone guidance. Neuron 43, 835–846 (2004). · DOI
  16. Mattila, P. K. & Lappalainen, P. Filopodia: molecular architecture and cellular functions. Nat. Rev. Mol. Cell Biol. 9, 446–454 (2008). · DOI
  17. Rajnicek, A. M., Foubister, L. E. & McCaig, C. D. Growth cone steering by a physiological electric field requires dynamic microtubules, microfilaments and Rac-mediated filopodial asymmetry. J. Cell. Sci. 119, 1736–1745 (2006). · DOI
  18. Zheng, J. Q., Wan, J. J. & Poo, M. M. Essential role of filopodia in chemotropic turning of nerve growth cone induced by a glutamate gradient. J. Neurosci. 16, 1140–1149 (1996). · DOI
  19. Lin, C. H., Espreafico, E. M., Mooseker, M. S. & Forscher, P. Myosin drives retrograde F-actin flow in neuronal growth cones. Neuron 16, 769–782 (1996). · DOI
  20. Medeiros, N. A., Burnette, D. T. & Forscher, P. Myosin II functions in actin-bundle turnover in neuronal growth cones. Nat. Cell Biol. 8, 215–226 (2006). · DOI
  21. Caswell, P. T. & Zech, T. Actin-based cell protrusion in a 3D matrix. Trends Cell Biol. 28, 823–834 (2018). · DOI
  22. Andrianantoandro, E. & Pollard, T. D. Mechanism of actin filament turnover by severing and nucleation at different concentrations of ADF/cofilin. Mol. Cell 24, 13–23 (2006). · DOI
  23. Cooper, J. A., Blum, J. D., Williams, R. C. & Pollard, T. D. Purification and characterization of actophorin, a new 15,000-dalton actin-binding protein from Acanthamoeba castellanii. J. Biol. Chem. 261, 477–485 (1986). · DOI
  24. Cruz, E. M. D. L. How cofilin severs an actin filament. Biophys. Rev. 1, 51–59 (2009). · DOI
  25. Wioland, H. et al. ADF/Cofilin accelerates actin dynamics by severing filaments and promoting their depolymerization at both ends. Curr. Biol. 27, 1956–1967.e7 (2017). · DOI
  26. Shekhar, S. & Carlier, M.-F. Enhanced depolymerization of actin filaments by ADF/cofilin and monomer funneling by capping protein cooperate to accelerate barbed-end growth. Curr. Biol. 27, 1990–1998.e5 (2017). · DOI
  27. Gungabissoon, R. A. & Bamburg, J. R. Regulation of growth cone actin dynamics by ADF/cofilin. J. Histochem. Cytochem. 51, 411–420 (2003). · DOI
  28. Tedeschi, A. et al. ADF/cofilin-mediated actin turnover promotes axon regeneration in the adult CNS. Neuron 103, 1073–1085.e6 (2019). · DOI
  29. Meberg, P. J. & Bamburg, J. R. Increase in neurite outgrowth mediated by overexpression of actin depolymerizing factor. J. Neurosci. 20, 2459–2469 (2000). · DOI
  30. Cruz, E. M. D. L. Cofilin binding to muscle and non-muscle actin filaments: isoform-dependent cooperative interactions. J. Mol. Biol. 346, 557–564 (2005). · DOI
  31. McGough, A., Pope, B., Chiu, W. & Weeds, A. Cofilin changes the twist of F-actin: implications for actin filament dynamics and cellular function. J. Cell Biol. 138, 771–781 (1997). · DOI
  32. Bobkov, A. A. et al. Cooperative effects of cofilin (ADF) on actin structure suggest allosteric mechanism of cofilin function. J. Mol. Biol. 356, 325–334 (2006). · DOI
  33. Cao, W., Goodarzi, J. P. & Cruz, E. M. D. L. Energetics and kinetics of cooperative cofilin–actin filament interactions. J. Mol. Biol. 361, 257–267 (2006). · DOI
  34. Elam, W. A. et al. Phosphomimetic S3D cofilin binds but only weakly severs actin filaments. J. Biol. Chem. 292, 19565–19579 (2017). · DOI
  35. Huehn, A. R. et al. Structures of cofilin-induced structural changes reveal local and asymmetric perturbations of actin filaments. Proc. Natl Acad. Sci. USA 117, 1478–1484 (2020). · DOI
  36. Bobkov, A. A. et al. Structural effects of cofilin on longitudinal contacts in F-actin. J. Mol. Biol. 323, 739–750 (2002). · DOI · Paper page
  37. Chou, S. Z. & Pollard, T. D. Mechanism of actin polymerization revealed by cryo-EM structures of actin filaments with three different bound nucleotides. Proc. Natl Acad. Sci. USA 116, 201807028 (2019). · DOI
  38. Pfaendtner, J., Cruz, E. M. D. L. & Voth, G. A. Actin filament remodeling by actin depolymerization factor/cofilin. Proc. Natl Acad. Sci. USA 107, 7299–7304 (2010). · DOI
  39. Tanaka, K. et al. Structural basis for cofilin binding and actin filament disassembly. Nat. Commun. 9, 1860–12 (2018). · DOI
  40. Galkin, V. E. et al. Remodeling of actin filaments by ADF/cofilin proteins. Proc. Natl Acad. Sci. USA 108, 20568–20572 (2011). · DOI
  41. Hocky, G. M., Sindelar, C. V., Cao, W., Voth, G. A. & Cruz, E. M. D. L. Structural basis of fast- and slow-severing actin-cofilactin boundaries. J. Biol. Chem. 296, 100337 (2021). · DOI
  42. McCullough, B. R. et al. Cofilin-linked changes in actin filament flexibility promote severing. Biophys. J. 101, 151–159 (2011). · DOI
  43. Ngo, K. X., Kodera, N., Katayama, E., Ando, T. & Uyeda, T. Q. P. Cofilin-induced unidirectional cooperative conformational changes in actin filaments revealed by high-speed atomic force microscopy. Elife 4, 393 (2015). · DOI
  44. Ono, S. Cofilin-induced structural changes in actin filaments stay local. Proc. Natl Acad. Sci. USA 117, 3349–3351 (2020). · DOI
  45. Suarez, C. et al. Cofilin tunes the nucleotide state of actin filaments and severs at bare and decorated segment boundaries. Curr. Biol. 21, 862–868 (2011). · DOI · Paper page
  46. Wioland, H., Jegou, A. & Romet-Lemonne, G. Quantitative variations with pH of actin depolymerizing factor/cofilin’s multiple actions on actin filaments. Biochemistry 58, 40–47 (2019). · DOI
  47. Hayden, S. M., Miller, P. S., Brauweiler, A. & Bamburg, J. R. Analysis of the interactions of actin depolymerizing factor with G- and F-actin. Biochemistry 32, 9994–10004 (1993). · DOI
  48. McCullough, B. R., Blanchoin, L., Martiel, J.-L. & Cruz, E. M. D. L. Cofilin increases the bending flexibility of actin filaments: implications for severing and cell mechanics. J. Mol. Biol. 381, 550–558 (2008). · DOI
  49. Prochniewicz, E., Janson, N., Thomas, D. D. & Cruz, E. M. D. L. Cofilin increases the torsional flexibility and dynamics of actin filaments. J. Mol. Biol. 353, 990–1000 (2005). · DOI
  50. Aramaki, S., Mayanagi, K., Jin, M., Aoyama, K. & Yasunaga, T. Filopodia formation by crosslinking of F-actin with fascin in two different binding manners. Cytoskeleton (Hoboken) 73, 365–374 (2016). · DOI
  51. Cohan, C. S., Welnhofer, E. A., Zhao, L., Matsumura, F. & Yamashiro, S. Role of the actin bundling protein fascin in growth cone morphogenesis: localization in filopodia and lamellipodia. Cell Motil. Cytoskeleton 48, 109–120 (2001). · DOI
  52. DeRosier, D. J. & Edds, K. T. Evidence for fascin cross-links between the actin filaments in coelomocyte filopodia. Exp. Cell Res. 126, 490–494 (1980). · DOI
  53. Jasnin, M. et al. Three-dimensional architecture of actin filaments in Listeria monocytogenes comet tails. Proc. Natl Acad. Sci. USA 110, 20521–20526 (2013). · DOI
  54. Vignjevic, D. et al. Role of fascin in filopodial protrusion. J. Cell Biol. 174, 863–875 (2006). · DOI
  55. Yang, S. et al. Molecular mechanism of fascin function in filopodial formation. J. Biol. Chem. 288, 274–284 (2013). · DOI
  56. Breitsprecher, D. et al. Cofilin cooperates with fascin to disassemble filopodial actin filaments. J. Cell. Sci. 124, 3305–3318 (2011). · DOI
  57. Elam, W. A., Kang, H. & Cruz, E. M. D. L. Competitive displacement of cofilin can promote actin filament severing. Biochem. Biophys. Res. Commun. 438, 728–731 (2013). · DOI
  58. Wiggan, O., Shaw, A. E., DeLuca, J. G. & Bamburg, J. R. ADF/cofilin regulates actomyosin assembly through competitive inhibition of myosin II binding to F-actin. Dev. Cell 22, 530–543 (2012). · DOI
  59. Goyal, P. et al. Cofilin oligomer formation occurs in vivo and is regulated by cofilin phosphorylation. PLoS One 8, e71769 (2013). · DOI
  60. Pfannstiel, J. et al. Human cofilin forms oligomers exhibiting actin bundling activity. J. Biol. Chem. 276, 49476–49484 (2001). · DOI
  61. Riedl, J. et al. Lifeact: a versatile marker to visualize F-actin. Nat. Methods 5, 605–607 (2008). · DOI · Paper page
  62. Bernstein, B. W., Shaw, A. E., Minamide, L. S., Pak, C. W. & Bamburg, J. R. Incorporation of cofilin into rods depends on disulfide intermolecular bonds: implications for actin regulation and neurodegenerative disease. J. Neurosci. 32, 6670–6681 (2012). · DOI
  63. Minamide, L. S., Striegl, A. M., Boyle, J. A., Meberg, P. J. & Bamburg, J. R. Neurodegenerative stimuli induce persistent ADF/cofilin-actin rods that disrupt distal neurite function. Nat. Cell Biol. 2, 628–636 (2000). · DOI
  64. Grintsevich, E. E. et al. F-actin dismantling through a Redox-driven synergy between mical and cofilin. Nat. Cell Biol. 18, 876–885 (2016). · DOI
  65. Grintsevich, E. E. et al. Catastrophic disassembly of actin filaments via Mical-mediated oxidation. Nat. Commun. 8, 2183 (2017). · DOI
  66. Chan, C., Beltzner, C. C. & Pollard, T. D. Cofilin dissociates Arp2/3 complex and branches from actin filaments. Curr. Biol. 19, 537–545 (2009). · DOI
  67. Huehn, A. et al. The actin filament twist changes abruptly at boundaries between bare and cofilin-decorated segments. J. Biol. Chem. 293, 5377–5383 (2018). · DOI
  68. Kremer, J. R., Mastronarde, D. N. & McIntosh, J. R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 116, 71–76 (1996). · DOI
  69. Fujii, T., Iwane, A. H., Yanagida, T. & Namba, K. Direct visualization of secondary structures of F-actin by electron cryomicroscopy. Nature 467, 724–728 (2010). · DOI
  70. Rigort, A. et al. Automated segmentation of electron tomograms for a quantitative description of actin filament networks. J. Struct. Biol. 177, 135–144 (2012). · DOI

Cited by

Sorted by citations

See all 45

Metadata

Authors
Hylton, R. K., Heebner, J. E., Grillo, M. A., & Swulius, M. T.
Affiliations
Pennsylvania State University (US)
Corresponding author
Swulius, Matthew T.
ORCID
Venue
Nature Communications
Publisher
Springer Science and Business Media LLC
Published
2022-05-04
Volume
13
Issue
1
Pages
2439-2439
ISSN
2041-1723 (ISSN-L)
DOI
10.1038/s41467-022-30116-x
PMC
PMC9068697
OpenAlex
W4225376724
Cited by
45
References
2
Citations by year
2026: 4 · 2025: 9 · 2024: 14 · 2023: 12 · 2022: 6
FWCI
5.47
Type
journal article
Language
en
Open access
Licence
Locations
  • Nature Communications (publishedVersion, cc-by)
  • PubMed Central (publishedVersion, cc-by)
  • PubMed (publishedVersion)
  • RePEc: Research Papers in Economics (submittedVersion)
  • DOAJ (DOAJ: Directory of Open Access Journals) (submittedVersion, cc-by-sa)
Funders
  • University of Pennsylvania
  • Pennsylvania State University (SCR_021178)
  • Penn State College of Medicine
Topics
  • Life Sciences › Biochemistry, Genetics and Molecular Biology › Cell Biology › Cellular Mechanics and Interactions
  • Life Sciences › Neuroscience › Cellular and Molecular Neuroscience › Axon Guidance and Neuronal Signaling
  • Life Sciences › Biochemistry, Genetics and Molecular Biology › Molecular Biology › Protein Kinase Regulation and GTPase Signaling
Keywords
cofilin, filopodia, growth cones, fascin, actin cross-linking, actin filament dynamics, cytoskeletal remodeling
MeSH terms
Actins, Actin Cytoskeleton, Pseudopodia, Growth Cones, Actin Depolymerizing Factors

About these checks

Registry, retraction, Cites and statistics results are computed, not judged, with one exception: a printed reference that no index lists is read once by a model before it counts as a citation error. Claims are extracted by a model and kept only when their quote appears verbatim in the text; each cited source is then checked, with a model’s help, for whether it says what the claim relies on. Those verdicts can be wrong — read the claim’s quote and the cited source before relying on one.

Statistics and arithmetic are recomputed, not judged.

Confidence bands are shown only for fields calibrated on our annotated test set. None is calibrated yet, so no band is shown.

What counts as a claim ›How we measure confidence ›How we check this ›

Are you an author, or do you see a problem with this page? Request a review