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References 1. Benson, N.C., Butt, O.H., Datta, R., Radoeva, P.D., Brainard, D.H., and Aguirre, G.K. (2012). The retinotopic organization of striate cortex is well predicted by surface topology. Curr. Biol. 22, 2081–2085. 2. Inouye, T. (1906). Visual disturbances following gunshot wounds of the cortical visual area by Tatsuji Inouye (translated by M. Glickstein & M. Fahle). Brain 123 (special suppl.), 1–100. 3. Holmes, G. (1918). Disturbances of visual orientation. Br. J. Ophthalmol. 2, 506–516. 4. Hinds, O., Polimeni, J.R., Rajendran, N., Balasubramanian, M., Amunts, K., Zilles, K., Schwartz, E.L., Fischl, B., and Triantafyllou, C. (2009). Locating the functional and anatomical boundaries of human primary visual cortex. Neuroimage 46, 915–922. 5. Schira, M.M., Tyler, C.W., Spehar, B., and Breakspear, M. (2010). Modeling magnification and anisotropy in the primate foveal confluence. PLoS Comput. Biol. 6, e1000651. 6. Van Essen, D.C. (1997). A tension-based theory of morphogenesis and compact wiring in the central nervous system. Nature 385, 313–318. 7. Gattass, R., Sousa, A.P., and Rosa, M.G. (1987). Visual topography of V1 in the Cebus monkey. J. Comp. Neurol. 259, 529–548. 8. Fritsches, K.A., and Rosa, M.G. (1996). Visuotopic organisation of striate cortex in the marmoset monkey (Callithrix jacchus). J. Comp. Neurol. 372, 264–282. 9. Rosa, M.G., Casagrande, V.A., Preuss, T., and Kaas, J.H. (1997). Visual field representation in
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striate and prestriate cortices of a prosimian primate (Galago garnetti). J. Neurophysiol. 77, 3193–3217. Gattass, R., Gross, C.G., and Sandell, J.H. (1981). Visual topography of V2 in the macaque. J. Comp. Neurol. 201, 519–539. Donoghue, M.J., and Rakic, P. (1999). Molecular evidence for the early specification of presumptive functional domains in the embryonic primate cerebral cortex. J. Neurosci. 19, 5967–5979. Rosa, M.G., and Tweedale, R. (2005). Brain maps, great and small: lessons from comparative studies of primate visual cortical organization. Phil. Trans. R. Soc. Lond. B 360, 665–691. Schira, M.M., Tyler, C.W., Breakspear, M., and Spehar, B. (2009). The foveal confluence in human visual cortex. J. Neurosci. 29, 9050–9058. Torab, K., Davis, T.S., Warren, D.J., House, P.A., Normann, R.A., and Greger, B. (2011). Multiple factors may influence the performance of a visual prosthesis based on intracortical microstimulation: nonhuman primate behavioural experimentation. J. Neural. Eng. 8, 035001. Schira, M.M., Wade, A.R., and Tyler, C.W. (2007). Two-dimensional mapping of the central and parafoveal visual field to human visual cortex. J. Neurophysiol. 97, 4284–4295. Chaplin, T.A., Yu, H.H., and Rosa, M.G.P. (2012). Representation of the visual field in the primary visual area of the marmoset monkey: magnification factors, point-image size and proportionality to retinal ganglion cell density.
Intracellular Transport: New Tools Provide Insights into Multi-motor Transport Teams of kinesin and dynein motors drive bidirectional transport of intracellular cargoes along the microtubule cytoskeleton. How do opposite-polarity motors interact to achieve targeted trafficking? A new study uses tools from synthetic biology to probe collective motor function. Adam G. Hendricks1, Alison E. Twelvetrees1,2, and Erika L.F. Holzbaur1 Many intracellular cargoes move bidirectionally along the microtubule cytoskeleton, transported by teams of kinesin and dynein motors [1]. Kinesin drives motility towards microtubule plus ends, while dynein moves towards the minus end. The collective function of motor teams allows cargoes to move bidirectionally over large distances. This long-range transport is vital for extended, polarized cells such as neurons. Indeed, defects in microtubule motors cause neurodegenerative disease, and impaired axonal transport has been identified in models of amyotrophic lateral sclerosis, and Alzheimer’s and
Huntington’s diseases [2]. Despite its fundamental importance, an understanding of how interactions among motors on a cargo modulate their function to achieve targeted trafficking in the cell remains poorly understood. Several mechanisms have been proposed to describe bidirectional transport. One hypothesis is that directional switches are the result of a regulatory event [1]. Alternatively, bidirectional motility may result from a stochastic tug-of-war between opposite polarity motors bound to the same cargo. In this case, switching is a consequence of the force-dependent dissociation kinetics of the motors, in the absence of regulation [3]. These mechanisms are not mutually exclusive, and both are likely to
J. Comp. Neurol. http://dx.doi.org/10.1002/ cne.23215. 17. Adams, D.L., and Horton, J.C. (2003). A precise retinotopic map of primate striate cortex generated from the representation of angioscotomas. J. Neurosci. 23, 3771–3789. 18. Polimeni, J.R., Balasubramanian, M., and Schwartz, E.L. (2006). Multi-area visuotopic map complexes in macaque striate and extra-striate cortex. Vision Res. 46, 3336–3359. 19. Van Essen, D.C., Drury, H.A., Dickson, J., Harwell, J., Hanlon, D., and Anderson, C.H. (2001). An integrated software suite for surface-based analyses of cerebral cortex. J. Am. Med. Inform. Assoc. 8, 443–459. 1Neuroscience Research Australia, Randwick, NSW, Australia & School of Psychology, University of Wollongong, Wollongong, NSW, Australia, 2Smith-Kettlewell Brain Imaging Center, Smith-Kettlewell Eye Research Institute, San Francisco, CA, USA, 3Department of Physiology and Monash Vision Group, Monash University, Melbourne, VIC, Australia. E-mail:
[email protected],
[email protected],
[email protected]
http://dx.doi.org/10.1016/j.cub.2012.11.003
contribute to the motility of intracellular cargoes [4]. Diverse approaches have been used to elucidate the mechanisms of multi-motor transport, ranging from in vitro reconstitution using purified motors bound to latex beads to high-resolution tracking of endogenous cargoes moving in the cell (Figure 1). In a recent paper published in Science, Derr et al. [5] implement a novel synthetic biology approach. The authors construct a scaffold using DNA origami — a technique that allows the creation of complex three-dimensional structures from DNA [6]. Motor proteins can be specifically attached to these artificial scaffolds via strands of complementary DNA, and in this way the number and type of motors can be tightly controlled. Derr et al. [5] use these scaffolds to examine motility by teams of dynein and kinesin motors. In agreement with previous work [7,8], the authors find that the run length of the cargoes increases with the number of motors, while velocity is largely unaffected [9]. They next examine teams of dynein and kinesin bound to the same cargo. Although the human kinesin-1 and yeast cytoplasmic dynein motors used in this study have similar unitary stall forces, a few dynein motors
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Figure 1. Techniques for studying motor function. (A) The synthetic DNA origami scaffold allows precise linkage of purified recombinant motor proteins, controlling both the number and the orientation of coupled motors [5]. (B) Optical trap methods for measuring motor forces use beads coupled to recombinant motor proteins. (C) TIRF microscopy can be used to study the motility of motor proteins in vitro. Either single molecules of fluorescent motor proteins can be imaged moving on immobilized filaments, or (as shown here) in the inverse filament gliding assay motor proteins are immobilized on glass while fluorophore-labelled filaments (actin or microtubules) are imaged as they move across the surface. (D) Cargoes/organelles can be purified or enriched from tissue or cell preparations. For example, compartments containing latex or magnetic beads internalized by phagocytosis can be purified. Frequently, preparations can be enriched for a specific organelle that is already labelled with a fluorophore. These endogenous cargoes co-purify with a cohort of bound factors that may or may not be regulating the activity of motors. Further, the nature of motor–cargo linkage is frequently unknown. (E) Recent advances in cellular studies of motor activity include the internalization of latex beads, which allow optical trap measurements of endogenous compartments as they move along microtubules. The motility of single molecules of fluorophore-labelled motor proteins or cargoes has also been studied in cells.
were observed to out-compete many kinesin motors. This result is surprising, but less so in the light of previous work using DNA scaffolds to link two kinesin
motors that also indicated that motility by teams of kinesin-1 is relatively insensitive to motor number [10]. Derr et al. [5] find that the number of
immotile cargoes increases as plus- and minus-end motors become more evenly matched, suggesting that the motors bound to these immotile cargoes are engaged in a static tug-of-war. In an elegant illustration of this point, immotile cargoes began to move when dynein motors were released from the scaffold through photo-cleavable linkages. This approach now provides a powerful technique to examine motor interactions in vitro, yet the motility of these motor-bound artificial scaffolds differs in significant ways from the observed transport of kinesin- and dynein-driven cargoes in cells. While artificial scaffolds move unidirectionally and are often stalled in a static tug-of-war, endogenous cargoes with opposing motors bound are motile and display frequent directional switches [11,12]. Vesicles isolated from mouse brain continue to move bidirectionally along microtubules in vitro, driven by a complement of stably-bound kinesin-1, kinesin-2, and dynein motors in the absence of cytosolic factors [4]. Quantitative analysis of active motors bound to endogenous cargoes suggests that bidirectional cargoes are driven by small teams of strong kinesin motors and large teams of relatively weak dynein motors operating at or near force balance [4,13–15]. One possible explanation for differences in the motility observed for artificial scaffolds and endogenous cargoes is that additional regulatory factors are required to reconstitute bidirectional motility. These regulatory factors may act to prevent the static tug-of-war observed for synthetic cargoes. Alternatively, the observed differences in motility might be due to the mechanochemistry of the specific motors involved. Derr et al. [5] paired human kinesin-1 with yeast cytoplasmic dynein. Consistent with its role in sliding microtubules along the cell cortex in vivo, yeast dynein is a slow (maximum velocity w85 nm/s), highly processive (run length w2 mm), and strong (unitary stall force w6 pN) motor [16,17]. In contrast, mammalian dynein is fast (maximum velocity w900 nm/s), less processive (run length w1 mm), and weak (unitary stall force w1 pN, although this value remains somewhat controversial) [13,18]. Another possible factor is the mechanical coupling between motors, which has been shown to affect the
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ability of multiple motors to work in teams [19]. Artificial scaffolds are expected to be more rigid than endogenous membranous vesicles; this rigidity may negatively affect motor coordination. These results highlight the need for the use of diverse approaches to understand the collective dynamics of motors. Engineered cargoes like those developed by Derr et al. [5] will be extremely useful in examining the effect of motor number and coupling and can be extended to include physiological motor complements and possible regulatory factors, including motor binding partners and effectors. In parallel, new techniques for imaging and manipulation, such as fast subpixel tracking [20] and optical trapping in living cells [14,15], can be used to examine the motility of endogenous cargoes with high resolution. Future work will allow a complete understanding of the collective dynamics of motor proteins in intracellular transport to emerge as techniques to design and manipulate minimal motor systems converge with high-resolution methods to examine transport in living cells. References 1. Welte, M.A. (2004). Bidirectional transport along microtubules. Curr. Biol. 14, R525–R537.
2. Hirokawa, N., Niwa, S., and Tanaka, Y. (2010). Molecular motors in neurons: transport mechanisms and roles in brain function, development, and disease. Neuron 68, 610–638. 3. Muller, M.J.I., Klumpp, S., and Lipowsky, R. (2008). Tug-of-war as a cooperative mechanism for bidirectional cargo transport by molecular motors. Proc. Natl. Acad. Sci. USA 105, 4609–4614. 4. Hendricks, A.G., Perlson, E., Ross, J.L., Schroeder, H.W., 3rd, Tokito, M., and Holzbaur, E.L.F. (2010). Motor coordination via a tug-of-war mechanism drives bidirectional vesicle transport. Curr. Biol. 20, 697–702. 5. Derr, N., Goodman, B., Jungmann, R., Leschziner, A., Shih, W., and Reck-Peterson, S. (2012). Tug-of-war in motor protein ensembles revealed with a programmable DNA origami scaffold. Science 338, 662–665. 6. Douglas, S., Dietz, H., Liedl, T., Hogberg, B., Graf, F., and Shih, W. (2009). Self-assembly of DNA into nanoscale three-dimensional shapes. Nature 459, 414–418. 7. Beeg, J., Klumpp, S., Dimova, R., Gracia, R.S., Unger, E., and Lipowsky, R. (2008). Transport of beads by several kinesin motors. Biophys. J. 94, 532–541. 8. Vershinin, M., Carter, B.C., Razafsky, D.S., King, S.J., and Gross, S.P. (2007). Multiple-motor based transport and its regulation by tau. Proc. Natl. Acad. Sci. USA 104, 87–92. 9. Howard, J., Hudspeth, A.J., and Vale, R.D. (1989). Movement of microtubules by single kinesin molecules. Nature 342, 154–158. 10. Jamison, D.K., Driver, J.W., Rogers, A.R., Constantinou, P.E., and Diehl, M.R. (2010). Two kinesins transport cargo primarily via the action of one motor: implications for intracellular transport. Biophys. J. 99, 2967–2977. 11. Kural, C., Kim, H., Syed, S., Goshima, G., Gelfand, V.I., and Selvin, P.R. (2005). Kinesin and dynein move a peroxisome in vivo: a tug-of-war or coordinated movement. Science 308, 1469–1472. 12. Levi, V., Serpinskaya, A.S., Gratton, E., and Gelfand, V.I. (2006). Organelle transport along
Cell Polarity: ParA-logs Gather around the Hub The chromosomal origin, chemotaxis arrays and flagellum of Vibrio cholerae congregate at the same pole of the cell. How? A recent study identifies a new pole-organizing protein, HubP, that recruits members of the ParA family of spatial regulators of subcellular structures to the pole. Clare L. Kirkpatrick and Patrick H. Viollier* With a volume smaller than that of many eukaryotic organelles, rod-shaped bacteria rely on polar organizers to maintain a high degree of cellular organization [1]. Such organizers can selectively direct factors (such as pili, flagellae or chemotaxis proteins) to the pole, while excluding others (for example, the cell division proteins) from the
extremities. Loss of such cell polarization results in misplacement and thus mis-inheritance of cellular structures, potentially compromising the integrity of the chromosome(s) and impairing other functions required for survival and fitness in the wild such as virulence and/or motility. Polar organizers are variable in primary structure and in function across different bacterial lineages. In the Gram-positive lineage, the coiled-coil domain protein DivIVA
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microtubules in Xenopus melanophores: Evidence for cooperation between multiple motors. Biophys. J. 90, 318–327. Soppina, V., Rai, A.K., Ramaiya, A.J., Barak, P., and Mallik, R. (2009). Tug-of-war between dissimilar teams of microtubule motors regu- lates transport and fission of endosomes. Proc. Natl. Acad. Sci. USA 106, 19381–19386. Hendricks, A.G., Holzbaur, E.L.F., and Goldman, Y.E. (2012). Force measurements on cargoes in living cells reveal collective dynamics of microtubule motors. Proc. Natl. Acad. Sci. USA 109, 18447–18452. Rai, A.K., Rai, A., Ramaiya, A.J., Jha, R., and Mallik, R. (2012). Molecular adaptations in dynein to generate large collective forces inside cells. Cell, in press. Reck-Peterson, S.L., Yildiz, A., Carter, A.P., Gennerich, A., Zhang, N., and Vale, R.D. (2006). Single-molecule analysis of dynein processivity and stepping behavior. Cell 126, 335–348. Gennerich, A., Carter, A.P., ReckPeterson, S.L., and Vale, R.D. (2007). Force-induced bidirectional stepping of cytoplasmic dynein. Cell 131, 952–965. Ross, J.L., Wallace, K., Shuman, H., Goldman, Y.E., and Holzbaur, E.L.F. (2006). Processive bidirectional motion of dynein-dynactin complexes in vitro. Nat. Cell Biol. 8, 562–570. Bieling, P., Telley, I., Piehler, J., and Surrey, T. (2008). Processive kinesins require loose mechanical coupling for efficient collective motility. EMBO Rep. 9, 1121–1127. Nan, X., Sims, P.A., and Xie, X.S. (2008). Organelle tracking in a living cell with microsecond time resolution and nanometer spatial precision. Chemphyschem 9, 707.
1University of Pennsylvania Perelman School of Medicine, Philadelphia, PA 19104, USA, 2Cancer Research UK London Research Institute, London WC2A 3LY, UK. E-mail:
[email protected]
http://dx.doi.org/10.1016/j.cub.2012.11.009
orchestrates polar activities by recruiting origin-binding proteins, cell division inhibitors, cell wall-modifying enzymes, and competence and secretion factors [2,3]. In the asymmetric Gram-negative alpha-proteobacterium Caulobacter crescentus, three unrelated coiled-coil motif-containing proteins TipN, PodJ and PopZ act as polar organizers to direct flagellar, pili and origin-binding proteins, respectively, to the newborn pole [4–9]. In addition, the muramidase homolog SpmX acts as an old-pole-specific localization factor for a developmental kinase [10]. The pathogenic Gram-negative gamma-proteobacterium Vibrio cholerae (causative agent of cholera) is another good example of polar organization, as it directs multiple cellular structures to the pole, namely, the origin of one (though not both) of its
