Encyclopedia of Signaling Molecules

2018 Edition
| Editors: Sangdun Choi

Myosin X

Reference work entry
DOI: https://doi.org/10.1007/978-3-319-67199-4_404

Synonyms

Historical Background

In 1994, myoX was first identified in a PCR screen designed to find novel  myosins in the inner ear (Solc et al. 1994). The full-length cDNA sequences of myoX were determined in 2000 in mouse (Yonezawa et al. 2000), human, and cow (Berg et al. 2000), and the molecular structure and ubiquitous expression in tissues were reported (Berg et al. 2000; Yonezawa et al. 2000). The unique localization pattern of myoX in cultured cells at the tips of filopodia and in lamellipodia was identified in the same reports (Berg et al. 2000; Yonezawa et al. 2000). Subsequently, in 2001, it was shown that the head domain of myoX binds actin and hydrolyzes ATP to induce movement and force (Homma et al. 2001). In 2002, myoX was shown to participate in intrafilopodial movement and filopodia formation (Berg and Cheney 2002), and in phagocytosis (Cox et al. 2002). After these breakthroughs, studies of myoX were expanded to many different aspects by precise biochemical analyses or by identifying binding partners critical for exploring its functions in cells.

Molecular Structure

The N-terminal domain of myoX functions as a motor domain: It binds actin and hydrolyzes ATP to produce movement and force (Homma et al. 2001). The motor domain is followed by a neck region that contains three IQ motifs, which bind calmodulin or calmodulin-like light chains (Berg et al. 2000). In epithelial cells, myoX binds calmodulin-like protein (CLP), an epithelial-specific light chain that is expressed during differentiation. CLP expression increases myoX protein levels by stabilizing the molecule (Bennett et al. 2007). A predicted coiled-coil segment is present at the C-terminus of the neck region (Berg et al. 2000); however, it has been suggested that this domain does not form a stable coiled-coil, but instead forms a stable α-helix (SAH) (Knight et al. 2005). The C-terminal end of the molecule is the tail domain, which consists of four different domains: PEST, PH, MTH4, and FERM (Fig. 1). Classes VII, X, XII, and XV myosins share a conserved structural feature in their tail domains – a myosin tail homology 4 (MTH4) domain followed by a band 4.1, Ezrin, Radixin, Moesin (FERM) domain. Together, the myosins containing these domains comprise the MTH4-FERM superclass of myosins (Berg et al. 2001).
Myosin X, Fig. 1

Schematic diagram of MyoX protein. The N-terminus of myoX functions as a motor domain. The motor domain is followed by a neck region. The predicted coiled-coil segment is present at the C-terminus of the neck region [22]; however, a recent study suggests that instead of forming a stable coiled-coil, this domain forms a stable α-helix (SAH) [23]. The C-terminus of the molecule, or tail, consists of four different domains: a PEST domain; three pleckstrin homology (PH) domains; a myosin tail homology 4 (MTH4) domain; and a band 4.1, Ezrin, Radixin, Moesin (FERM) domain

In addition to this full-length myoX, brain expresses a shorter form that lacks a motor domain (Sousa et al. 2006). This “headless” myoX might have a possible role as a natural dominant-negative regulator to suppress the function of full-length myoX in neurons.

Intrafilopodial Motility

One of the most striking properties of myoX is its forward and rearward movements within filopodia (Berg and Cheney 2002). Recently, the velocity of the forward movement was calculated as ∼600 nm/s in living cells using TIRF microscopy (Kerber et al. 2009), similar to the 340–780 nm/s reported for movement of individual myoX molecules on artificial actin bundles (Nagy et al. 2008). The rearward movement of myoX in filopodia is slow at 10–20 nm/s (Berg and Cheney 2002), presumably the rate of retrograde actin flow in filopodia. In these studies, only dimerized myoX showed intrafilopodial motility, suggesting that dimer formation of myoX is necessary for its proper movement in cells.

A forced dimer construct of tail-less myoX was first reported to preferentially select bundled actin for motility and showed poor processivity on single filaments in vitro (Nagy et al. 2008). The selectivity of actin tracks (single filaments or bundled) by myoX is controversial because a recent report shows the robust processivity of myoX on individual actin filaments (Sun et al. 2010). Sun et al. also report that myoX moves processively in a hand-over-hand manner with a left-hand helical walking path using single-molecule fluorescence techniques such as polTIRF, FIONA, and Parallax.

Filopodia Formation

MyoX has an important role in the formation of filopodia (Berg et al. 2000; Tokuo and Ikebe 2004; Bohil et al. 2006; Tokuo et al. 2007; Zhu et al. 2007), but the mechanism of filopodia induction by myoX is still largely unknown. Using a regulated dimerization technique (Tokuo et al. 2007), it was shown that the motor activity of myoX is itself critical for the initiation of filopodia formation. The regulated dimerization system (Ariad Pharmaceuticals) was based on the human FK506-binding protein variant (FKBP) and its small molecular ligands. An EGFP-tagged myoX construct was produced by fusing FKBP to the C-terminal end of the SAH domain (EGFP-M10MoIQ3SAH-FKBP, Fig. 2a). If two FKBPs are present, the dimer-inducing drug, AP20187, binds to both FKBPs, thus creating a dimer of the target molecule (Fig. 2b). After addition of dimerizer, EGFP-M10MoIQ3SAH-FKBP induced filopodia in transfected cells (Fig. 2b). These results support the following model (Fig. 3): (a) MyoX is present as a dimer and monomer in cells. Monomeric (single-headed) myoX does not localize at the edge of lamellipodia. (b) Once dimerization occurs, myoX moves to the tip of the actin filaments, presumably as a result of its ability to walk toward the barbed ends of actin filaments. (c) The tips move laterally along the leading edge with actin filaments, and the mechanical activity of myoX plays a role in this process. (d) The clustering of myoX causes convergence of the actin filaments into parallel bundles, thus producing the base of the filopodia. This hypothetical model is elicited from the observation that dimerized tail-less myoX induces microspikes or short unstable filopodia which is thought to be an initial step in filopodia formation. On the other hand, deletion of the MTH4-FERM region abolishes the formation of stable and elongated dorsal filopodia by myoX (Bohil et al. 2006). From these results, it is plausible that monomer-to-dimer transition of the motor and neck region controls initiation, and the tail region controls elongation and stabilization of filopodia formation.
Myosin X, Fig. 2

Dimerization of myoX is critical for filopodia formation. NIH3T3 cells were transfected with GFP-M10MoIQ3SAH-FKBP (green) and replated on fibronectin-coated cover slips with (b) or without (a) the dimerizer AP20187. Red: rhodamine-phalloidin staining

Myosin X, Fig. 3

The filopodia initiation model by dimerization of myoX. (a) MyoX is present as a dimer and a monomer in cells. Monomeric (single-headed) myoX does not localize at the edge of lamellipodia. (b) Once the dimer is produced, myoX moves to and concentrates at the tip of actin filaments presumably due to its ability to walk toward the barbed end of actin filaments. (c) The tips move laterally along the leading edge with actin filaments, and myoX motor activity plays a role in this process. (d) The lateral movement of myoX causes the barbed end of the actin filaments to converge, thus producing the base of filopodia, where actin polymerization might induce formation of parallel actin bundles

TIRF microscopy was used to clarify the role of the tail domain in the mechanism of myoX-induced filopodia formation (Watanabe et al. 2010). MyoX was recruited to discrete sites at the leading edge where it assembles with exponential kinetics before filopodia extension. MyoX-induced filopodia showed repeated extension–retraction cycles with each extension of 2.4 μm, which was critical to produce long filopodia. FERM domain–deleted myoX moved to the tip as in wild type, but it was transported toward the cell body during filopodia retraction, did not undergo multiple extension–retraction cycles, and failed to produce long filopodia. Deletion of the FERM domain did not change movement at the single-molecule level with the same velocity of approximately 600 nm/s as wild type, suggesting that the myoX in filopodia moves without interacting with the attached membrane via the FERM domain. These results suggest that the interaction of myoX and substrate-engaged integrin is necessary for phased elongation.

Regulating Molecules and Other Functions

Recent studies implicate myoX in a variety of cellular functions through its interactions with other molecules:

PIP3: In leukocytes, myoX localizes to phagocytic cups with phosphatidylinositol-3,4,5-triphosphate (PIP3) in a PI3K-dependent manner. Expression in macrophages of a tail domain of myoX inhibits phagocytosis (Cox et al. 2002). MyoX might provide a molecular link between PI3K and pseudopod extension during phagocytosis.

Ena/VASP: MyoX and VASP bind in vitro and in vivo, colocalize at the tips of filopodia, move together in filopodia, and there is a correlation between the length of filopodia and the concentration of VASP/myoX at the tips of filopodia (Tokuo and Ikebe 2004). These results suggest that myoX transports VASP to the tips of filopodia to support elongation by actin incorporation.

Microtubules: Through its MTH4-FERM domain, MyoX associates with microtubules. Expression of the tail domain or microinjection of anti-myoX antibodies disrupts nuclear anchoring, spindle assembly, and spindle-F-actin association in Xenopus laevis (Weber et al. 2004). These results indicate that during meiosis, myoX has a critical role in integrating the F-actin and microtubule cytoskeletons. MyoX is also essential for mitotic-spindle function (Woolner et al. 2008). Interaction with microtubules is also important in osteoclast function (McMichael et al. 2010). MyoX suppression by RNAi or overexpression of dominant-negative myoX (MTH4-FERM) leads to decreased sealing zone perimeter, motility, and resorptive capacity of osteoclasts. These results suggest that myoX plays a role in osteoclast attachment and podosome positioning by direct linkage of actin to the microtubule network.

Integrins: The FERM domain of myoX interacts with an NPXY motif within the cytoplasmic domain of β-integrin (Zhang et al. 2004). Knockdown of myoX results in decreased integrin-mediated cell adhesion to the extracellular matrix, and myoX is responsible for localization of integrins in filopodia. Localization of integrin at filopodial tips and filopodia elongation did not occur with myoX mutants deficient in integrin binding or with a β-integrin mutant deficient in myoX binding. These results suggest that binding of myoX to integrins allows tethering of the filopodial actin filaments to the extracellular matrix for stabilizing the structure of filopodia and elongation.

BMP6: In endothelial cells, myoX is a target gene of bone morphogenetic protein (BMP), and myoX colocalizes with the BMP6 receptor, ALK6, in a BMP6-dependent fashion. Other data indicate that myoX is required to guide endothelial migration toward BMP6 gradients via the regulation of filopodial function and amplification of BMP signals (Pi et al. 2007).

Netrin receptor: The FERM domain of myoX interacts with the cytoplasmic tail of the netrin receptors, DCC (deleted in colorectal cancer) and neogenin (Zhu et al. 2007). Netrins regulate axon path-finding, which is essential for proper wiring in the brain. Cortical explants derived from mouse embryos expressing dominant-negative myoX exhibit reduced neurite outgrowth in response to netrin-1. Inhibition of myoX in embryos causes impaired commissural neuronal axon projections in chicken brain. These results indicate that myoX regulates axon outgrowth and guidance in response to netrins (Zhu et al. 2007).

Using Xenopus as a model system to study the function of myoX in neurons, two reports show that knock-down of myoX expression results in retarded migration of cranial neural crest cells (Hwang et al. 2009; Nie et al. 2009). These results suggest that myoX has an essential function in neuronal development in vertebrates.

VE–cadherin: MyoX is directly associated with the VE–cadherin complex via a FERM domain. It colocalizes and moves synchronously with filopodial VE–cadherin. Expression of the FERM domain blocks the transportation of VE–cadherin along actin fibers, resulting in an almost total depletion of VE–cadherin at the cell edge. VE–cadherin trafficking along filopodia by myoX may be a pre-requisite for cell–cell junction formation in endothelial cells (Almagro et al. 2010).

Summary

MyoX functions both as a molecular transporter and a cytoskeletal regulator in cells.

References

  1. Almagro S, Durmort C, Chervin-Petinot A, Heyraud S, Dubois M, Lambert O, et al. The motor protein myosin-X transports VE-cadherin along filopodia to allow the formation of early endothelial cell-cell contacts. Mol Cell Biol. 2010;30(7):1703–17.PubMedPubMedCentralCrossRefGoogle Scholar
  2. Bennett RD, Mauer AS, Strehler EE. Calmodulin-like protein increases filopodia-dependent cell motility via up-regulation of myosin-10. J Biol Chem. 2007;282(5):3205–12.PubMedCrossRefGoogle Scholar
  3. Berg JS, Cheney RE. Myosin-X is an unconventional myosin that undergoes intrafilopodial motility. Nat Cell Biol. 2002;4(3):246–50.PubMedCrossRefGoogle Scholar
  4. Berg JS, Derfler BH, Pennisi CM, Corey DP, Cheney RE. Myosin-X, a novel myosin with pleckstrin homology domains, associates with regions of dynamic actin. J Cell Sci. 2000;113(Pt 19):3439–51.PubMedGoogle Scholar
  5. Berg JS, Powell BC, Cheney RE. A millennial myosin census. Mol Biol Cell. 2001;12(4):780–94.PubMedPubMedCentralCrossRefGoogle Scholar
  6. Bohil AB, Robertson BW, Cheney RE. Myosin-X is a molecular motor that functions in filopodia formation. Proc Natl Acad Sci USA. 2006;103(33):12411–6.PubMedPubMedCentralCrossRefGoogle Scholar
  7. Cox D, Berg JS, Cammer M, Chinegwundoh JO, Dale BM, Cheney RE, et al. Myosin X is a downstream effector of PI(3)K during phagocytosis. Nat Cell Biol. 2002;4(7):469–77.PubMedGoogle Scholar
  8. Homma K, Saito J, Ikebe R, Ikebe M. Motor function and regulation of myosin X. J Biol Chem. 2001;276(36):34348–54.PubMedCrossRefGoogle Scholar
  9. Hwang YS, Luo T, Xu Y, Sargent TD. Myosin-X is required for cranial neural crest cell migration in Xenopus laevis. Dev Dyn. 2009;238(10):2522–9.PubMedPubMedCentralCrossRefGoogle Scholar
  10. Kerber ML, Jacobs DT, Campagnola L, Dunn BD, Yin T, Sousa AD, et al. A novel form of motility in filopodia revealed by imaging myosin-X at the single-molecule level. Curr Biol. 2009;19(11):967–73.PubMedPubMedCentralCrossRefGoogle Scholar
  11. Knight PJ, Thirumurugan K, Xu Y, Wang F, Kalverda AP, Stafford 3rd WF, et al. The predicted coiled-coil domain of myosin 10 forms a novel elongated domain that lengthens the head. J Biol Chem. 2005;280(41):34702–8.PubMedCrossRefGoogle Scholar
  12. McMichael BK, Cheney RE, Lee BS. Myosin X regulates sealing zone patterning in osteoclasts through linkage of podosomes and microtubules. J Biol Chem. 2010;285(13):9506–15.PubMedPubMedCentralCrossRefGoogle Scholar
  13. Nagy S, Ricca BL, Norstrom MF, Courson DS, Brawley CM, Smithback PA, et al. A myosin motor that selects bundled actin for motility. Proc Natl Acad Sci USA. 2008;105(28):9616–20.PubMedPubMedCentralCrossRefGoogle Scholar
  14. Nie S, Kee Y, Bronner-Fraser M. Myosin-X is critical for migratory ability of Xenopus cranial neural crest cells. Dev Biol. 2009;335(1):132–42.PubMedPubMedCentralCrossRefGoogle Scholar
  15. Pi X, Ren R, Kelley R, Zhang C, Moser M, Bohil AB, et al. Sequential roles for myosin-X in BMP6-dependent filopodial extension, migration, and activation of BMP receptors. J Cell Biol. 2007;179(7):1569–82.PubMedPubMedCentralCrossRefGoogle Scholar
  16. Solc CK, Derfler BH, Duyk GM, Corey DP. Molecular cloning of myosins from the bullfrog saccular macula: a candidate for the hair cell adaptation motor. Aud Neurosci. 1994;1:63–75.Google Scholar
  17. Sousa AD, Berg JS, Robertson BW, Meeker RB, Cheney RE. Myo10 in brain: developmental regulation, identification of a headless isoform and dynamics in neurons. J Cell Sci. 2006;119(Pt 1):184–94.PubMedCrossRefGoogle Scholar
  18. Sun Y, Sato O, Ruhnow F, Arsenault ME, Ikebe M, Goldman YE. Single-molecule stepping and structural dynamics of myosin X. Nat Struct Mol Biol. 2010;17(4):485–91.PubMedPubMedCentralCrossRefGoogle Scholar
  19. Tokuo H, Ikebe M. Myosin X transports Mena/VASP to the tip of filopodia. Biochem Biophys Res Commun. 2004;319(1):214–20.PubMedCrossRefGoogle Scholar
  20. Tokuo H, Mabuchi K, Ikebe M. The motor activity of myosin-X promotes actin fiber convergence at the cell periphery to initiate filopodia formation. J Cell Biol. 2007;179(2):229–38.PubMedPubMedCentralCrossRefGoogle Scholar
  21. Watanabe TM, Tokuo H, Gonda K, Higuchi H, Ikebe M. Myosin-X induces filopodia by multiple elongation mechanism. J Biol Chem. 2010;285:19605–14.PubMedPubMedCentralCrossRefGoogle Scholar
  22. Weber KL, Sokac AM, Berg JS, Cheney RE, Bement WM. A microtubule-binding myosin required for nuclear anchoring and spindle assembly. Nature. 2004;431(7006):325–9.PubMedCrossRefGoogle Scholar
  23. Woolner S, O’Brien LL, Wiese C, Bement WM. Myosin-10 and actin filaments are essential for mitotic spindle function. J Cell Biol. 2008;182(1):77–88.PubMedPubMedCentralCrossRefGoogle Scholar
  24. Yonezawa S, Kimura A, Koshiba S, Masaki S, Ono T, Hanai A, et al. Mouse myosin X: molecular architecture and tissue expression as revealed by northern blot and in situ hybridization analyses. Biochem Biophys Res Commun. 2000;271(2):526–33.PubMedCrossRefGoogle Scholar
  25. Zhang H, Berg JS, Li Z, Wang Y, Lang P, Sousa AD, et al. Myosin-X provides a motor-based link between integrins and the cytoskeleton. Nat Cell Biol. 2004;6(6):523–31.PubMedCrossRefGoogle Scholar
  26. Zhu XJ, Wang CZ, Dai PG, Xie Y, Song NN, Liu Y, et al. Myosin X regulates netrin receptors and functions in axonal path-finding. Nat Cell Biol. 2007;9(2):184–92.PubMedCrossRefGoogle Scholar

Copyright information

© Springer International Publishing AG 2018

Authors and Affiliations

  1. 1.Boston Biomedical Research InstituteWatertownUSA