Key Points
-
Vertebrate myosin Va is structurally and kinetically designed to be able to move processively along actin filaments as a single molecule and is thus well suited to be a cargo transporter. The key features are the presence of two motor domains and kinetics with a high duty ratio.
-
Not all class V myosins are structurally or kinetically similar to vertebrate myosin Va. Some do not have high duty ratios and are not two headed. These class V myosins must adopt other strategies to move cargo, such as having multiple motors bound to the cargo.
-
Budding yeast and plants build actin cytoskeletons that are suitable for long range class V myosin-dependent cargo transport. Indeed, the yeast class V myosins Myo2 and Myo4 drive most, if not all, organelle transport in this organism.
-
The organization of actin within the cortex of vertebrate cells is largely anisotropic, so the class V myosin-dependent transport of organelles in these cells, which in most cases will follow the long-range transport of the organelle on microtubules to the cell periphery, is likely to exhibit minimal directional persistence and be very local.
-
Two recent and compelling examples of class V myosin-dependent organelle transport have been identified within the dendritic spines of neurons, one involving recycling endosomes, and the other involving the endoplasmic reticulum.
-
The 'acid test' as to whether a class V myosin actually moves an organelle in vivo is to show that the organelle moves more slowly when the cell expresses a 'slower' version of the class V myosin.
Abstract
Cells use molecular motors, such as myosins, to move, position and segregate their organelles. Class V myosins possess biochemical and structural properties that should make them ideal actin-based cargo transporters. Indeed, studies show that class V myosins function as cargo transporters in yeast, moving a range of organelles, such as the vacuole, peroxisomes and secretory vesicles. There is also increasing evidence in vertebrate cells that class V myosins not only tether organelles to actin but also can serve as short-range, point-to-point organelle transporters, usually following long-range, microtubule-dependent organelle transport.
This is a preview of subscription content, access via your institution
Access options
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Odronitz, F. & Kollmar, M. Drawing the tree of eukaryotic life based on the analysis of 2269 manually annotated myosins from 328 species. Genome Biol. 8, R196 (2007).
Sellers, J. R. Myosins (Oxford University Press, 1999).
Sakamoto, T. et al. Neck length and processivity of myosin V. J. Biol. Chem. 278, 29201–29207 (2003).
Howard, J. & Spudich, J. A. Is the lever arm of myosin a molecular elastic element? Proc. Natl Acad. Sci. USA 93, 4462–4464 (1996).
Warshaw, D. M. et al. The light chain binding domain of expressed smooth muscle heavy meromyosin acts as a mechanical lever. J. Biol. Chem. 275, 37167–37172 (2000).
Ruff, C., Furch, M., Brenner, B., Manstein, D. J. & Meyhofer, E. Single-molecule tracking of myosins with genetically engineered amplifier domains. Nature Struct. Biol. 8, 226–229 (2001).
Krendel, M. & Mooseker, M. S. Myosins: tails (and heads) of functional diversity. Physiology (Bethesda) 20, 239–251 (2005).
Berg, J. S., Powell, B. C. & Cheney, R. E. A millennial myosin census. Mol. Biol. Cell 12, 780–794 (2001).
Mehta, A. D. et al. Myosin-V is a processive actin-based motor. Nature 400, 590–593 (1999). Shows, for the first time, that class V myosins move processively along actin filaments with 36 nm step sizes, using optical trapping nanometry.
Sakamoto, T., Amitani, I., Yokota, E. & Ando, T. Direct observation of processive movement by individual myosin V molecules. Biochem. Biophys. Res. Commun. 272, 586–590 (2000).
Cheney, R. E. et al. Brain myosin-V is a two-headed unconventional myosin with motor activity. Cell 75, 13–23 (1993).
Warshaw, D. M. et al. Differential labeling of myosin V heads with quantum dots allows direct visualization of hand-over-hand processivity. Biophys. J. 88, L30–L32 (2005).
Churchman, L. S., Okten, Z., Rock, R. S., Dawson, J. F. & Spudich, J. A. Single molecule high-resolution colocalization of Cy3 and Cy5 attached to macromolecules measures intramolecular distances through time. Proc. Natl Acad. Sci. USA 102, 1419–1423 (2005).
Walker, M. L. et al. Two-headed binding of a processive myosin to F-actin. Nature 405, 804–807 (2000). Presents electron microscopic images of negatively stained class V myosin molecules trapped in the process of moving along actin, demonstrating that the two heads are bound 36 nm apart.
Veigel, C., Wang, F., Bartoo, M. L., Sellers, J. R. & Molloy, J. E. The gated gait of the precessive molecular motor, myosin V. Nature Cell Biol. 4, 59–65 (2001).
Rief, M. et al. Myosin-V stepping kinetics: a molecular model for processivity. Proc. Natl Acad. Sci. USA 97, 9482–9486 (2000).
Yildiz, A. et al. Myosin V walks hand-over-hand: single fluorophore imaging with 1.5-nm localization. Science 300, 2061–2065 (2003). Describes the use of super-resolution light microscopy to follow the movement of one head of myosin V as it moved along an actin filament bound to a coverslip surface, demonstrating a hand-over-hand walking mechanism.
Snyder, G. E., Sakamoto, T., Hammer, J. A. III, Sellers, J. R. & Selvin, P. R. Nanometer localization of single green fluorescent proteins: evidence that myosin V walks hand-over-hand via telemark configuration. Biophys. J. 87, 1776–1783 (2004).
Kodera, N., Yamamoto, D., Ishikawa, R. & Ando, T. Video imaging of walking myosin V by high-speed atomic force microscopy. Nature 468, 72–76 (2010).
Nagy, A., Piszczek, G. & Sellers, J. R. Extensibility of the extended tail domain of processive and nonprocessive myosin V molecules. Biophys. J. 97, 3123–3131 (2009).
Schilstra, M. J. & Martin, S. R. An elastically tethered viscous load imposes a regular gait on the motion of myosin-V. Simulation of the effect of transient force relaxation on a stochastic process. J. R. Soc. Interface 3, 153–165 (2005).
Wu, X. S. et al. Identification of an organelle receptor for myosin-Va. Nature Cell Biol. 4, 271–278 (2002). Uses various approaches, including the characterization of melanocytes isolated from three mouse coat-colour mutants, to show that the RAB GTPase RAB27A and its effector protein melanophilin serve as the melanosome receptor for myosin Va. See also references 24–26.
Wu, X., Wang, F., Rao, K., Sellers, J. R. & Hammer, J. A. III. Rab27a is an essential component of melanosome receptor for myosin Va. Mol. Biol. Cell 13, 1735–1749 (2002).
Fukuda, M., Kuroda, T. S. & Mikoshiba, K. Slac2-a/melanophilin, the missing link between Rab27 and myosin Va: implications of a tripartite protein complex for melanosome transport. J. Biol. Chem. 277, 12432–12436 (2002).
Strom, M., Hume, A. N., Tarafder, A. K., Barkagianni, E. & Seabra, M. C. A family of Rab27-binding proteins. Melanophilin links Rab27a and myosin Va function in melanosome transport. J. Biol. Chem. 277, 25423–25430 (2002).
Nagashima, K. et al. Melanophilin directly links Rab27a and myosin Va through its distinct coiled-coil regions. FEBS Lett. 517, 233–238 (2002).
Wagner, W., Fodor, E., Ginsburg, A. & Hammer, J. A. III. The binding of DYNLL2 to myosin Va requires alternatively spliced exon B and stabilizes a portion of the myosin's coiled-coil domain. Biochemistry 45, 11564–11577 (2006).
Hodi, Z. et al. Alternatively spliced exon B of myosin Va is essential for binding the tail-associated light chain shared by dynein. Biochemistry 45, 12582–12595 (2006).
Schroeder, H. W. III, Mitchell, C., Shuman, H., Holzbaur, E. L. & Goldman, Y. E. Motor number controls cargo switching at actin-microtubule intersections in vitro. Curr. Biol. 20, 687–696 (2010).
Ali, M. Y. et al. Myosin Va maneuvers through actin intersections and diffuses along microtubules. Proc. Natl Acad. Sci. USA 104, 4332–4336 (2007).
De La Cruz, E. M., Wells, A. L., Rosenfeld, S. S., Ostap, E. M. & Sweeney, H. L. The kinetic mechanism of myosin V. Proc. Natl Acad. Sci. USA 96, 13726–13731 (1999).
Veigel, C., Schmitz, S., Wang, F. & Sellers, J. R. Load-dependent kinetics of myosin-V can explain its high processivity. Nature Cell Biol. 7, 861–869 (2005).
Rosenfeld, S. S. & Sweeney, H. L. A model of myosin V processivity. J. Biol. Chem. 279, 40100–40111 (2004).
Purcell, T. J., Sweeney, H. L. & Spudich, J. A. A force-dependent state controls the coordination of processive myosin V. Proc. Natl Acad. Sci. USA 102, 13873–13878 (2005).
Forgacs, E. et al. Kinetics of ADP dissociation from the trail and lead heads of actomyosin V following the power stroke. J. Biol. Chem. 283, 766–773 (2008).
Sakamoto, T., Webb, M. R., Forgacs, E., White, H. D. & Sellers, J. R. Direct observation of the mechanochemical coupling in myosin Va during processive movement. Nature 455, 128–132 (2008).
Thirumurugan, K., Sakamoto, T., Hammer, J. A. III, Sellers, J. R. & Knight, P. J. The cargo-binding domain regulates structure and activity of myosin 5. Nature 442, 212–215 (2006).
Liu, J., Taylor, D. W., Krementsova, E. B., Trybus, K. M. & Taylor, K. A. Three-dimensional structure of the myosin V inhibited state by cryoelectron tomography. Nature 442, 208–211 (2006). References 37 and 38 reveal the conformation of the folded, quiescent state of class V myosin. Reference 37 uses single-particle averaging of negatively stained class V myosin molecules in solution. Reference 38 uses cryo-electron microscopy of class V myosin molecules bound to a lipid monolayer.
Wang, F. et al. Regulated conformation of myosin V. J. Biol. Chem. 279, 2333–2336 (2004).
Sato, O., Li, X. D. & Ikebe, M. Myosin Va becomes a low duty ratio motor in the inhibited form. J. Biol. Chem. 282, 13228–13239 (2007).
Lu, H., Krementsova, E. B. & Trybus, K. M. Regulation of myosin V processivity by calcium at the single molecule level. J. Biol. Chem. 281, 31987–31994 (2006).
Trybus, K. M. et al. Effect of calcium on calmodulin bound to the IQ motifs of myosin V. J. Biol. Chem. 282, 23316–23325 (2007).
Nguyen, H. & Higuchi, H. Motility of myosin V regulated by the dissociation of single calmodulin. Nature Struct. Mol. Biol. 12, 127–132 (2005).
Wu, X., Sakamoto, T., Zhang, F., Sellers, J. R. & Hammer, J. A. III. In vitro reconstitution of a transport complex containing Rab27a, melanophilin and myosin Va. FEBS Lett. 580, 5863–5868 (2006).
Li, X. D., Ikebe, R. & Ikebe, M. Activation of myosin Va function by melanophilin, a specific docking partner of myosin Va. J. Biol. Chem. 280, 17815–17822 (2005).
Watanabe, S., Mabuchi, K., Ikebe, R. & Ikebe, M. Mechanoenzymatic characterization of human myosin Vb. Biochemistry 45, 2729–2738 (2006).
Takagi, Y. et al. Human myosin Vc is a low duty ratio, non-processive molecular motor. J. Biol. Chem. 283, 8527–8537 (2008).
Toth, J., Kovacs, M., Wang, F., Nyitray, L. & Sellers, J. R. Myosin V from Drosophila reveals diversity of motor mechanisms within the myosin V family. J. Biol. Chem. 280, 30594–30603 (2005).
Watanabe, S. et al. Human myosin Vc is a low duty ratio nonprocessive motor. J. Biol. Chem. 283, 10581–10592 (2008).
Taft, M. H. et al. Dictyostelium myosin-5b is a conditional processive motor. J. Biol. Chem. 283, 26902–26910 (2008).
Dunn, B. D., Sakamoto, T., Hong, M. S., Sellers, J. R. & Takizawa, P. A. Myo4p is a monomeric myosin with motility uniquely adapted to transport mRNA. J. Cell Biol. 178, 1193–1206 (2007).
Hodges, A. R., Bookwalter, C. S., Krementsova, E. B. & Trybus, K. M. A nonprocessive class V myosin drives cargo processively when a kinesin-related protein is a passenger. Curr. Biol. 19, 2121–2125 (2009).
Hodges, A. R., Krementsova, E. B. & Trybus, K. M. She3p binds to the rod of yeast myosin V and prevents it from dimerizing, forming a single-headed motor complex. J. Biol. Chem. 283, 6906–6914 (2008).
Pruyne, D., Legesse-Miller, A., Gao, L., Dong, Y. & Bretscher, A. Mechanisms of polarized growth and organelle segregation in yeast. Annu. Rev. Cell Dev. Biol. 20, 559–591 (2004).
Weisman, L. S. Yeast vacuole inheritance and dynamics. Annu. Rev. Genet. 37, 435–460 (2003).
Fagarasanu, A., Mast, F. D., Knoblach, B. & Rachubinski, R. A. Molecular mechanisms of organelle inheritance: lessons from peroxisomes in yeast. Nature Rev. Mol. Cell Biol. 11, 644–654 (2010).
Schott, D. H., Collins, R. N. & Bretscher, A. Secretory vesicles transport velocity in living cells depends on the myosin-V lever arm. J. Cell Biol. 156, 35–39 (2002). By showing that the speed of secretory vesicle transport decreases in yeast Myo2-null cells complemented with 'slower' versions of Myo2 (that is, the first application of the acid test referred to in this Review), these authors provide unequivocal evidence that this class V myosin drives secretory vesicle transport.
Lipatova, Z. et al. Direct interaction between a myosin V motor and the Rab GTPases Ypt31/32 is required for polarized secretion. Mol. Biol. Cell 19, 4177–4187 (2008).
Santiago-Tirado, F. H., Legesse-Miller, A., Schott, D. & Bretscher, A. PI4P and Rab inputs collaborate in myosin-V-dependent transport of secretory compartments in yeast. Dev. Cell 20, 47–59 (2011).
Mizuno-Yamasaki, E., Medkova, M., Coleman, J. & Novick, P. Phosphatidylinositol 4-phosphate controls both membrane recruitment and a regulatory switch of the Rab GEF Sec2p. Dev. Cell 18, 828–840 (2010). References 59 and 60 reveal the key components of the receptor for Myo2 on yeast secretory vesicles. Both also elucidate key aspects of the regulation of Myo2 recruitment, which involves the membrane lipid PtdIns4P and sequential association with two different Rab GTPases.
Casavola, E. C. et al. Ypt32p and Mlc1p bind within the vesicle binding region of the class V myosin Myo2p globular tail domain. Mol. Microbiol. 67, 1051–1066 (2008).
Graham, T. R. & Burd, C. G. Coordination of Golgi functions by phosphatidylinositol 4-kinases. Trends Cell Biol. 21, 113–121 (2011).
Rossanese, O. W. et al. A role for actin, Cdc1p, and Myo2p in the inheritance of late Golgi elements in Saccharomyces cerevisiae. J. Cell Biol. 153, 47–62 (2001).
Arai, S., Noda, Y., Kainuma, S., Wada, I. & Yoda, K. Ypt11 functions in bud-directed transport of the Golgi by linking Myo2 to the coatomer subunit Ret2. Curr. Biol. 18, 987–991 (2008).
Hill, K. L., Catlett, N. L. & Weisman, L. S. Actin and myosin function in directed vacuole movement during cell division in Saccharomyces cerevisae. J. Cell Biol. 135, 1535–1549 (1996).
Ishikawa, K. et al. Identification of an organelle-specific myosin V receptor. J. Cell Biol. 160, 887–897 (2003).
Tang, F. et al. Regulated degradation of a class V myosin receptor directs movement of the yeast vacuole. Nature 422, 87–92 (2003). References 66 and 67 define the receptor for Myo2 on the surface of the yeast vacuole. Reference 67 also shows that the cell cycle-regulated degradation of a component of the vacuole receptor for Myo2 facilitates the correct deposition of the vacuole in the bud.
Peng, Y. & Weisman, L. S. The cyclin-dependent kinase Cdk1 directly regulates vacuole inheritance. Dev. Cell 15, 478–485 (2008).
Fagarasanu, A., Fagarasanu, M., Eitzen, G. A., Aitchison, J. D. & Rachubinski, R. A. The peroxisomal membrane protein Inp2p is the peroxisome-specific receptor for the myosin V motor Myo2p of Saccharomyces cerevisiae. Dev. Cell 10, 587–600 (2006).
Chang, J. et al. Pex3 peroxisome biogenesis proteins function in peroxisome inheritance as class V myosin receptors. J. Cell Biol. 187, 233–246 (2009).
Fagarasanu, A. et al. Myosin-driven peroxisome partitioning in S. cerevisiae. J. Cell Biol. 186, 541–554 (2009).
Fagarasanu, M., Fagarasanu, A., Tam, Y. Y., Aitchison, J. D. & Rachubinski, R. A. Inp1p is a peroxisomal membrane protein required for peroxisome inheritance in Saccharomyces cerevisiae. J. Cell Biol. 169, 765–775 (2005).
Bobola, N., Jansen, R. P., Shin, T. H. & Nasmyth, K. Asymmetric accumulation of ASH1p in postanaphase nuclei depends on a myosin and restricts yeast mating-type switching to mother cells. Cell 84, 699–709 (1996).
Takizawa, P. A., Sil, A., Swedlow, J. R., Herskowitz, I. & Vale, R. D. Actin-dependent localization of an RNA encoding a cell-fate determinant in yeast. Nature 389, 90–93 (1997).
Shepard, K. A. et al. Widespread cytoplasmic mRNA transport in yeast: identification of 22 bud-localized transcripts using DNA microarray analysis. Proc. Natl Acad. Sci. USA 100, 11429–11434 (2003).
Bertrand, E. et al. Localization of ASH1 mRNA particles in living yeast. Mol. Cell 2, 437–445 (1998).
Chung, S. & Takizawa, P. A. Multiple Myo4 motors enhance ASH1 mRNA transport in Saccharomyces cerevisiae. J. Cell Biol. 189, 755–767 (2010). Shows how the clustering of multiple monomeric Myo4 molecules by the self-association of the adaptor proteins that link it to mRNA allows this non-processive motor to drive processive mRNA transport in vivo.
Long, R. M., Gu, W., Lorimer, E., Singer, R. H. & Chartrand, P. She2p is a novel RNA-binding protein that recruits the Myo4p–She3p complex to ASH1 mRNA. EMBO J. 19, 6592–6601 (2000).
Kruse, C. et al. Ribonucleoprotein-dependent localization of the yeast class V myosin Myo4p. J. Cell Biol. 159, 971–982 (2002).
Yoshimura, A. et al. Myosin-Va facilitates the accumulation of mRNA/protein complex in dendritic spines. Curr. Biol. 16, 2345–2351 (2006).
Krauss, J., Lopez de, Q. S., Nusslein-Volhard, C. & Ephrussi, A. Myosin-V regulates Oskar mRNA localization in the Drosophila oocyte. Curr. Biol. 19, 1058–1063 (2009).
Estrada, P. et al. Myo4p and She3p are required for cortical ER inheritance in Saccharomyces cerevisiae. J. Cell Biol. 163, 1255–1266 (2003).
Fehrenbacher, K. L., Davis, D., Wu, M., Boldogh, I. & Pon, L. A. Endoplasmic reticulum dynamics, inheritance, and cytoskeletal interactions in budding yeast. Mol. Biol. Cell 13, 854–865 (2002).
Schmid, M., Jaedicke, A., Du, T. G. & Jansen, R. P. Coordination of endoplasmic reticulum and mRNA localization to the yeast bud. Curr. Biol. 16, 1538–1543 (2006).
Jung, G., Titus, M. A. & Hammer, J. A. III. The Dictyostelium type V myosin MyoJ is responsible for the cortical association and motility of contractile vacuole membranes. J. Cell Biol. 186, 555–570 (2009).
Satoh, A. K., Li, B. X., Xia, H. & Ready, D. F. Calcium-activated Myosin V closes the Drosophila pupil. Curr. Biol. 18, 951–955 (2008).
Li, B. X., Satoh, A. K. & Ready, D. F. Myosin V, Rab11, and dRip11 direct apical secretion and cellular morphogenesis in developing Drosophila photoreceptors. J. Cell Biol. 177, 659–669 (2007).
Woolner, S. & Bement, W. M. Unconventional myosins acting unconventionally. Trends Cell Biol. 19, 245–252 (2009).
Langford, G. M. Actin- and microtubule-dependent organelle motors: interrelationships between the two motility systems. Curr. Opin. Cell Biol. 7, 82–88 (1995).
Nascimento, A. A., Roland, J. T. & Gelfand, V. I. Pigment cells: a model for the study of organelle transport. Annu. Rev. Cell Dev. Biol. 19, 469–491 (2003).
Provance, D. W., James, T. L. & Mercer, J. A. Melanophilin, the product of the leaden locus, is required for targeting of myosin-Va to melanosomes. Traffic. 3, 124–132 (2002).
Wu, X., Bowers, B., Rao, K., Wei, Q. & Hammer, J. A. III. Visualization of melanosome dynamics within wild-type and dilute melanocytes suggests a paradigm for myosin V function in vivo. J. Cell Biol. 143, 1899–1918 (1998). Shows, by comparing melanosome distribution and dynamics in wild-type versus dilute (myosin Va-null) melanocytes, that melanosome positioning is driven by a cooperation between long-range, microtubule-dependent melanosome transport and myosin Va-dependent melanosome capture (and possibly local movement) in the cell periphery (the cooperative-capture model).
Chabrillat, M. L. et al. Rab8 regulates the actin-based movement of melanosomes. Mol. Biol. Cell 16, 1640–1650 (2005).
Rodionov, V. I., Hope, A. J., Svitkina, T. M. & Borisy, G. G. Functional coordination of microtubule-based and actin-based motility in melanophores. Curr. Biol. 8, 165–168 (1998).
Rogers, S. L. & Gelfand, V. I. Myosin cooperates with microtubule motors during organelle transport in melanophores. Curr. Biol. 8, 161–164 (1998). References 94 and 95 show that, in amphibian pigment cells, microtubule-based and actomyosin-V-based melanosome transport cooperate to drive the correct dispersion of the organelles required for the darkening of the animal.
Rogers, S. L. et al. Regulation of melanosome movement in the cell cycle by reversible association with myosin V. J. Cell Biol. 146, 1265–1276 (1999).
Semenova, I. et al. Actin dynamics is essential for myosin-based transport of membrane organelles. Curr. Biol. 18, 1581–1586 (2008).
Kural, C. et al. Tracking melanosomes inside a cell to study molecular motors and their interaction. Proc. Natl Acad. Sci. USA 104, 5378–5382 (2007).
Lapierre, L. A. et al. Myosin Vb is associated with plasma membrane recycling systems. Mol. Biol. Cell 12, 1843–1857 (2001).
Hales, C. M., Vaerman, J. P. & Goldenring, J. R. Rab11 family interacting protein 2 associates with Myosin Vb and regulates plasma membrane recycling. J. Biol. Chem. 277, 50415–50421 (2002).
Lapierre, L. A. & Goldenring, J. R. Interactions of myosin Vb with Rab11 family members and cargoes traversing the plasma membrane recycling system. Methods Enzymol. 403, 715–723 (2005).
Akhmanova, A. & Hammer, J. A. III. Linking molecular motors to membrane cargo. Curr. Opin. Cell Biol. 22, 479–487 (2010).
Roland, J. T. et al. Rab GTPase-Myo5B complexes control membrane recycling and epithelial polarization. Proc. Natl Acad. Sci. USA 108, 2789–2794 (2011).
Muller, T. et al. MYO5B mutations cause microvillus inclusion disease and disrupt epithelial cell polarity. Nature Genet. 40, 1163–1165 (2008).
Park, M., Penick, E. C., Edwards, J. G., Kauer, J. A. & Ehlers, M. D. Recycling endosomes supply AMPA receptors for LTP. Science 305, 1972–1975 (2004).
Wang, Z. et al. Myosin Vb mobilizes recycling endosomes and AMPA receptors for postsynaptic plasticity. Cell 135, 535–548 (2008). Provides multiple lines of support for the idea that myosin Vb transports recycling endosomes into the dendritic spines of hippocampal neurons in response to strong spine stimulation. These endosomes then serve as a source of AMPA receptors for insertion into the postsynaptic membrane to drive LTP.
Sellers, J. R., Thirumurugan, K., Sakamoto, T., Hammer, J. A. III & Knight, P. J. Calcium and cargoes as regulators of myosin 5a activity. Biochem. Biophys. Res. Commun. 369, 176–181 (2008).
Correia, S. S. et al. Motor protein-dependent transport of AMPA receptors into spines during long-term potentiation. Nature Neurosci. 11, 457–466 (2008).
Schnell, E. & Nicoll, R. A. Hippocampal synaptic transmission and plasticity are preserved in myosin Va mutant mice. J. Neurophysiol. 85, 1498–1501 (2001).
Petralia, R. S. et al. Glutamate receptor targeting in the postsynaptic spine involves mechanisms that are independent of myosin Va. Eur. J. Neurosci. 13, 1722–1732 (2001).
Takagishi, Y. et al. The dilute-lethal (dl) gene attacks a Ca2+ store in the dendritic spine of Purkinje cells in mice. Neurosci. Lett. 215, 169–172 (1996).
Miyata, M. et al. Local calcium release in dendritic spines required for long-term synaptic depression. Neuron 28, 233–244 (2000).
Wagner, W., Brenowitz, S. D. & Hammer, J. A. III. Myosin-Va transports the endoplasmic reticulum into the dendritic spines of Purkinje neurons. Nature Cell Biol. 13, 40–48 (2011). Provides, by using (among other things) the acid test referred to in this Review, clear evidence that myosin Va serves as a point-to-point transporter to move tubules of ER into the dendritic spines of cerebellar Purkinje neurons, which is required for the local Ca2+ transients that drive LTD.
Higashi-Fujime, S. & Nakamura, A. Cell and molecular biology of the fastest myosins. Int. Rev. Cell Mol. Biol. 276, 301–347 (2009).
Verchot-Lubicz, J. & Goldstein, R. E. Cytoplasmic streaming enables the distribution of molecules and vesicles in large plant cells. Protoplasma 240, 99–107 (2010).
Pollard, T. D. & Borisy, G. G. Cellular motility driven by assembly and disassembly of actin filaments. Cell 112, 453–465 (2003).
Desnos, C. et al. Myosin Va mediates docking of secretory granules at the plasma membrane. J. Neurosci. 27, 10636–10645 (2007).
Kogel, T. et al. Distinct roles of myosin Va in membrane remodeling and exocytosis of secretory granules. Traffic. 11, 637–650 (2010).
Nelson, S. R., Ali, M. Y., Trybus, K. M. & Warshaw, D. M. Random walk of processive, quantum dot-labeled myosin Va molecules within the actin cortex of COS-7 cells. Biophys. J. 97, 509–518 (2009). Describes, using quantum dot-labelled myosin Va in living cells, short-range movements of myosin Va in areas of the cell cortex dominated by actin filaments arranged anisotropically.
Pierobon, P. et al. Velocity, processivity, and individual steps of single myosin V molecules in live cells. Biophys. J. 96, 4268–4275 (2009).
Small, J. V. Dicing with dogma: de-branching the lamellipodium. Trends Cell Biol. 20, 628–633 (2010).
Kogel, T., Bittins, C. M., Rudolf, R. & Gerdes, H. H. Versatile roles for myosin Va in dense core vesicle biogenesis and function. Biochem. Soc. Trans. 38, 199–204 (2010).
Desnos, C., Huet, S. & Darchen, F. 'Should I stay or should I go?': myosin V function in organelle trafficking. Biol. Cell 99, 411–423 (2007).
Coudrier, E. Myosins in melanocytes: to move or not to move? Pigment Cell Res. 20, 153–160 (2007).
Loubery, S. & Coudrier, E. Myosins in the secretory pathway: tethers or transporters? Cell. Mol. Life Sci. 65, 2790–2800 (2008).
Provance, D. W. Jr et al. Myosin-Vb functions as a dynamic tether for peripheral endocytic compartments during transferrin trafficking. BMC. Cell Biol. 9, 44 (2008).
Watanabe, M. et al. Myosin-Va regulates exocytosis through the submicromolar Ca2+-dependent binding of syntaxin-1A. Mol. Biol. Cell 16, 4519–4530 (2005).
Fukuda, M. & Itoh, T. Slac2-a/melanophilin contains multiple PEST-like sequences that are highly sensitive to proteolysis. J. Biol. Chem. 279, 22314–22321 (2004).
Figueiredo, A. C. et al. Rab3GEP is the non-redundant guanine nucleotide exchange factor for Rab27a in melanocytes. J. Biol. Chem. 283, 23209–23216 (2008).
Itoh, T. & Fukuda, M. Identification of EPI64 as a GTPase-activating protein specific for Rab27A. J. Biol. Chem. 281, 31823–31831 (2006).
Kukimoto-Niino, M. et al. Structural basis for the exclusive specificity of Slac2-a/melanophilin for the Rab27 GTPases. Structure. 16, 1478–1490 (2008).
Fukuda, M. & Kuroda, T. S. Missense mutations in the globular tail of myosin-Va in dilute mice partially impair binding of Slac2-a/melanophilin. J. Cell Sci. 117, 583–591 (2004).
Karcher, R. L. et al. Cell cycle regulation of myosin-V by calcium/calmodulin-dependent protein kinase II. Science 293, 1317–1320 (2001).
Kuroda, T. S., Ariga, H. & Fukuda, M. The actin-binding domain of Slac2-a/melanophilin is required for melanosome distribution in melanocytes. Mol. Cell. Biol. 23, 5245–5255 (2003).
Wu, X. S., Tsan, G. L. & Hammer, J. A. III. Melanophilin and myosin Va track the microtubule plus end on EB1. J. Cell Biol. 171, 201–207 (2005).
Hume, A. N., Ushakov, D. S., Tarafder, A. K., Ferenczi, M. A. & Seabra, M. C. Rab27a and MyoVa are the primary Mlph interactors regulating melanosome transport in melanocytes. J. Cell Sci. 120, 3111–3122 (2007).
Hume, A. N., Tarafder, A. K., Ramalho, J. S., Sviderskaya, E. V. & Seabra, M. C. A coiled-coil domain of melanophilin is essential for myosin Va recruitment and melanosome transport in melanocytes. Mol. Biol. Cell 17, 4720–4735 (2006).
Krementsov, D. N., Krementsova, E. B. & Trybus, K. M. Myosin V: regulation by calcium, calmodulin, and the tail domain. J. Cell Biol. 164, 877–886 (2004).
Li, X. D., Mabuchi, K., Ikebe, R. & Ikebe, M. Ca2+-induced activation of ATPase activity of myosin Va is accompanied with a large conformational change. Biochem. Biophys. Res. Commun. 315, 538–545 (2004).
Jedd, G., Mulholland, J. & Segev, N. Two new Ypt GTPases are required for exit from the yeast trans-Golgi compartment. J. Cell Biol. 137, 563–580 (1997).
Jedd, G., Richardson, C., Litt, R. & Segev, N. The Ypt1 GTPase is essential for the first two steps of the yeast secretory pathway. J. Cell Biol. 131, 583–590 (1995).
Pashkova, N., Jin, Y., Ramaswamy, S. & Weisman, L. S. Structural basis for myosin V discrimination between distinct cargoes. EMBO J. 25, 693–700 (2006). Provides the first crystal structure for the GTD of a class V myosin, as well as insight into the mechanisms by which Myo2's GTD can bind multiple organelle-specific receptors. Opens the door to in-depth structure–function analyses of other class V myosin GTDs by homology modelling.
Mallik, R. & Gross, S. P. Molecular motors: strategies to get along. Curr. Biol. 14, R971–R982 (2004).
Hotulainen, P. & Hoogenraad, C. C. Actin in dendritic spines: connecting dynamics to function. J. Cell Biol. 189, 619–629 (2010).
Korobova, F. & Svitkina, T. Molecular architecture of synaptic actin cytoskeleton in hippocampal neurons reveals a mechanism of dendritic spine morphogenesis. Mol. Biol. Cell 21, 165–176 (2010).
Frost, N. A., Kerr, J. M., Lu, H. E. & Blanpied, T. A. A network of networks: cytoskeletal control of compartmentalized function within dendritic spines. Curr. Opin. Neurobiol. 20, 578–587 (2010).
Acknowledgements
We thank M. Peckham for drafting figure 1. For space reasons, we have not included many examples in which the evidence of class V myosin-dependant organelle transport, although suggestive, is not yet robust. We apologize to the authors of those studies for not citing their work.
Author information
Authors and Affiliations
Ethics declarations
Competing interests
The authors declare no competing financial interests.
Related links
Glossary
- Power stroke
-
The distance that the lever arm of myosin moves in a single event.
- Brownian diffusion
-
The random, thermally driven motion of small objects in a fluid or gas.
- Optical trapping
-
A technique that uses focused laser light in a light microscope to capture and manipulate objects with dielectric constants different from water. Sophisticated traps can be used to measure the mechanical properties of single molecules of motor proteins, such as the class V myosins.
- Super-resolution light microscopy
-
Techniques that use Gaussian fits to the point spread function of light emitted from a single fluorophore to determine its position to within a few nanometres, which is significantly smaller than the diffraction limit.
- Total internal reflection fluorescence
-
(TIRF). TIRF microscopy provides improved signal to noise ratios at the interface between media with differing refractive indices (such as coverslip–water), where fluorescence is excited by an evanescent field of light created when the incident light is totally internally reflected. It is ideal for visualizing the movement of single motor proteins.
- High-speed atomic force microscope
-
(High-speed AFM). A modified AFM that can acquire images at a high rate from samples in aqueous solution, allowing the dynamic imaging of single molecules as they perform their tasks.
- Myosin II
-
Also called conventional muscle myosin. The first myosin type to be discovered and the most conspicuous of the myosin superfamily. It is responsible for skeletal muscle contraction in muscle cells.
- Duty ratio
-
The fractional time that a myosin spends in a state of high affinity for actin during an ATP hydrolysis cycle.
- Melanosome
-
The pigment-producing organelle found in pigment-producing cells, such as melanocytes.
- RAB switch
-
The sequential interaction of an effector protein with two different RAB GTPases.
- Exocyst
-
An eight-member protein complex that is required for vesicle docking and polarized exocytosis and is conserved from yeast to mammals.
- Myristoylation
-
A protein post-translational modification in which the 14-carbon saturated fatty acid myristic acid is covalently attached to an amino-terminal Gly residue.
- Palmitoylation
-
A protein post-translational modification in which the 16-carbon saturated fatty acid palmitic acid is covalently attached to a Cys residue.
- PEST site
-
A sequence within proteins that serves as a site for cleavage by the Ca2+- dependent protease calpain.
- Recycling endosome
-
A generic, centrally located membrane compartment that receives endocytosed membrane receptors and recycles them back to the plasma membrane.
- Long-term potentiation
-
(LTP). A form of synaptic plasticity that is thought to underlie memory formation, in which synapse use leads to long-term strengthening of the synapse.
- AMPA receptors
-
The major excitatory ionotropic Glu receptors found in neurons. Their name comes from their ability to be activated by the artificial Glu analogue AMPA (α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid).
- Long-term depression
-
(LTD). A form of synaptic plasticity that is thought to underlie memory formation, in which synapse use leads to long-term weakening of the synapse.
- Anisotropic
-
A term referring to a complete lack of uniformity in orientation.
- Filopodia
-
Thin, dynamic, cellular extensions containing actin filaments aligned in parallel with their barbed ends pointing towards the tip. They are often found in growth cones and at the leading edge of migrating cells.
Rights and permissions
About this article
Cite this article
Hammer, J., Sellers, J. Walking to work: roles for class V myosins as cargo transporters. Nat Rev Mol Cell Biol 13, 13–26 (2012). https://doi.org/10.1038/nrm3248
Published:
Issue Date:
DOI: https://doi.org/10.1038/nrm3248
This article is cited by
-
Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons
Neuroscience Bulletin (2024)
-
Removal of a partial genomic duplication restores synaptic transmission and behavior in the MyosinVA mutant mouse Flailer
BMC Biology (2023)
-
Vesicle trafficking and vesicle fusion: mechanisms, biological functions, and their implications for potential disease therapy
Molecular Biomedicine (2022)
-
Autophagosome–lysosome fusion is facilitated by plectin-stabilized actin and keratin 8 during macroautophagic process
Cellular and Molecular Life Sciences (2022)
-
Let it go: mechanisms that detach myosin V from the yeast vacuole
Current Genetics (2021)