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Myosin Motors Drive Long Range Alignment of Actin Filaments2

  • Tariq Butt
    Affiliations
    From the Departments of Life Sciences, LUMS School of Science and Engineering,Sector U-DHA, Lahore 54792, Pakistan
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  • Tabish Mufti
    Affiliations
    Departments of Computer Science, LUMS School of Science and Engineering,Sector U-DHA, Lahore 54792, Pakistan and
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  • Ahmad Humayun
    Affiliations
    Departments of Computer Science, LUMS School of Science and Engineering,Sector U-DHA, Lahore 54792, Pakistan and
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  • Peter B. Rosenthal
    Affiliations
    the Division of Physical Biochemistry, MRC National Institute for Medical Research, Mill Hill, London NW7 1AA, United Kingdom
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  • Sohaib Khan
    Affiliations
    Departments of Computer Science, LUMS School of Science and Engineering,Sector U-DHA, Lahore 54792, Pakistan and
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  • Shahid Khan
    Correspondence
    To whom correspondence may be addressed
    Affiliations
    From the Departments of Life Sciences, LUMS School of Science and Engineering,Sector U-DHA, Lahore 54792, Pakistan
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  • Justin E. Molloy
    Correspondence
    To whom correspondence may be addressed: MRC NIMR, The Ridgeway, Mill Hill, London NW7 1AA, United Kingdom
    Affiliations
    the Division of Physical Biochemistry, MRC National Institute for Medical Research, Mill Hill, London NW7 1AA, United Kingdom
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  • Author Footnotes
    * The work was supported by a Higher Education Commission-British Council Link grant in “Bionanotechnology” (to Sh. K.), Lahore University of Management Sciences, School of Science and Engineering, Computer Science Department for research assistant stipends (to A. H. and T. M.), and a Medical Research Council, UK, grant-in-aid (to J. E. M. and P. B. R.).
    The on-line version of this article (available at http://www.jbc.org) contains supplemental movies.
Open AccessPublished:November 24, 2009DOI:https://doi.org/10.1074/jbc.M109.044792
      The bulk alignment of actin filament sliding movement, powered by randomly oriented myosin molecules, has been observed and studied using an in vitro motility assay. The well established, actin filament gliding assay is a minimal experimental system for studying actomyosin motility. Here, we show that when the assay is performed at densities of actin filaments approaching those found in living cells, filament gliding takes up a preferred orientation. The oriented patterns of movement that we have observed extend over a length scale of 10–100 μm, similar to the size of a mammalian cell. We studied the process of filament alignment and found that it depends critically upon filament length and density. We developed a simple quantitative measure of filament sliding orientation and this enabled us to follow the time course of alignment and the formation and disappearance of oriented domains. Domains of oriented filaments formed spontaneously and were separated by distinct boundaries. The pattern of the domain structures changed on the time scale of several seconds and the collision of neighboring domains led to emergence of new patterns. Our results indicate that actin filament crowding may play an important role in structuring the leading edge of migrating cells. Filament alignment due to near-neighbor mechanical interactions can propagate over a length scale of several microns; much greater than the size of individual filaments and analogous to a log drive. Self-alignment of actin filaments may make an important contribution to cell polarity and provide a mechanism by which cell migration direction responds to chemical cues.

      Introduction

      In vitro motility assays (
      • Yanagida T.
      • Nakase M.
      • Nishiyama K.
      • Oosawa F.
      ,
      • Kron S.J.
      • Spudich J.A.
      ) provide a defined biochemical system to characterize the mechanochemistry of motor proteins and cytoskeletal polymer dynamics (
      • Mizuno D.
      • Tardin C.
      • Schmidt C.F.
      • Mackintosh F.C.
      ). The actin filament gliding assay (
      • Kron S.J.
      • Spudich J.A.
      ) in particular, enables the movement of individual actin filaments to be observed, by fluorescence video microscopy. Rhodamine-phalloidin-labeled filaments are visualized as they move on a microscope coverslip surface that has been coated with randomly oriented myosin molecules. In the absence of Mg-ATP the filaments bind tightly to the surface, but when Mg-ATP is added, the filaments start sliding over the surface. Sequential video images, captured by computer, allow filament positions to be tracked in time using image analysis software so that the speed and direction of filament sliding can be determined. Under standard conditions the movement of an individual filament shows directed motion over a length scale of about 5–10 μm but after the filament has traveled though a contour distance of 100 μm its direction is completely randomized. So within a typical field of view a number of filaments will be seen traveling in a variety of directions. However, by using microfabrication techniques the microscope coverslip surface can be patterned so that filament sliding becomes confined to narrow, linear tracks (
      • Bunk R.
      • Klinth J.
      • Montelius L.
      • Nicholls I.A.
      • Omling P.
      • Tågerud S.
      • Månsson A.
      ). In living cells, myosins power motile processes in a crowded environment far removed from the conditions that are often used in in vitro assays. In particular, within the highly ordered sarcomeres of skeletal muscle cells, actin and myosin filaments are precisely oriented along the length of the cell. This means that external forces generated by muscle act along the axis of its fibers. Muscle cells are highly specialized to produce large external forces but every cell in the body contains myosin and actin and of the 39 human myosin genes most of them encode for so-called non-muscle myosins. Myosins are involved in a wide variety of cell motilities including vesicle trafficking and cell migration. The molecular mechanism of force generation by actomyosin has been extensively studied and the current view is as follows. Myosin binds actin ATP hydrolysis products (ADP and Pi) bound at its catalytic site; as the products are released the light chain binding region of the molecule swings through an angle of about 60° in a process known as the “power stroke”; the power stroke generates a relative sliding motion between the tail of the myosin and its attachment point with actin between 5 and 30 nm and a force of around 2 to 5 piconewtons; once the power stroke is complete a fresh ATP molecule binds to myosin, which causes it to detach from actin and reset its conformation (known as the “recovery stroke”); it can then undergo further cycles of interaction with actin as above. The time myosin spends tightly bound to actin compared with the total ATPase cycle time is called the “duty cycle ratio.” Fast acting myosins like those found in muscle typically have a low duty cycle ratio, whereas processive motors like myosin V have a high duty ratio (
      • Howard J.
      ). The myosins known to be involved in cell migration (myosin I, IIa, IIb, and IIc) (
      • Bresnick A.R.
      ) contribute to extension or retraction of the lamellipodium by driving the relative flow of cortical actin filaments past the plasma membrane and cell-substrate adhesions (
      • Diefenbach T.J.
      • Latham V.M.
      • Yimlamai D.
      • Liu C.A.
      • Herman I.M.
      • Jay D.G.
      ). The myosin II proteins localize by immunofluorescence in a punctate pattern at cell edges and their cellular role is under active study (
      • Wylie S.R.
      • Chantler P.D.
      ). Several other unconventional myosins such as myosin V, VI, VII, IX, and X may also play critical roles in formation and maintenance of the lamellipodial structure (
      • Small J.V.
      • Stradal T.
      • Vignal E.
      • Rottner K.
      ). The cortical actin forms a dense meshwork at the leading edge of the cell and the local actin concentration reaches 100 μm. This meshwork is stabilized and remodeled by various actin-related and actin-binding proteins (
      • Pollard T.D.
      ). Actin polymerization-depolymerization and actin bulk flow have been proposed to drive lamellipodial movements, in addition to or instead of myosin (
      • Verkhovsky A.B.
      • Svitkina T.M.
      • Borisy G.G.
      ,
      • Theriot J.A.
      • Mitchison T.J.
      ,
      • Pollard T.D.
      • Borisy G.G.
      ). Finally, accessory protein complexes involved in intracellular signaling couple actin dynamics to physiological function (
      • Takenawa T.
      • Miki H.
      ,
      • Symons M.H.
      • Mitchison T.J.
      ,
      • Ridley A.J.
      • Schwartz M.A.
      • Burridge K.
      • Firtel R.A.
      • Ginsberg M.H.
      • Borisy G.
      • Parsons J.T.
      • Horwitz A.R.
      ,
      • Hall A.
      ) and either activate or inhibit actin filament turnover and/or actomyosin interactions.
      Concentrated F-actin solutions undergo gel-sol phase transitions (
      • Wagner B.
      • Tharmann R.
      • Haase I.
      • Fischer M.
      • Bausch A.R.
      ) and the rheology and viscoelastic behavior (
      • Wiggins C.H.
      • Riveline D.
      • Ott A.
      • Goldstein R.E.
      ) has been well studied. It is known that actin filaments become aligned by flow and the shear modulus is strongly dependent on filament length (
      • Janmey P.A.
      • Hvidt S.
      • Käs J.
      • Lerche D.
      • Maggs A.
      • Sackmann E.
      • Schliwa M.
      • Stossel T.P.
      ). It is of interest whether filament alignment might arise spontaneously within the cell cortex due to shear forces and filament movement generated by actomyosin interactions. We have addressed this issue using an in vitro actin filament gliding assay in which actin filament density approaches that found in the cell cortex. We used computerized analysis of filament motility under a variety of experimental conditions in which filament movement is actively driven by surface-attached skeletal muscle, myosin II molecules.
      A number of algorithms have been described to track actin filament movement in in vitro motility assays (
      • Sheetz M.P.
      • Block S.M.
      • Spudich J.A.
      ,
      • Work S.S.
      • Warshaw D.M.
      ,
      • Marston S.B.
      • Fraser I.D.
      • Bing W.
      • Roper G.
      ,
      • Uttenweiler D.
      • Veigel C.
      • Steubing R.
      • Götz C.
      • Mann S.
      • Haussecker H.
      • Jähne B.
      • Fink R.H.
      ) and only a limited number of these methods have the capability to simultaneously measure the motion of many particles captured on video. This ability is essential for the present study, as we wish to investigate cooperative movements of many filaments simultaneously. We developed measures of the orientation of motile filament tracks at the single and population level. We used an established centroiding method (
      • Mashanov G.I.
      • Molloy J.E.
      ) and found this worked well for short filaments that could be treated as rigid rods, but worked less well for long, highly curved, filaments. So, we developed a new algorithm that works from skeletonized filament outlines and this approach enabled us to quantify filament velocity and direction at high filament densities. Our results give insight into how cortical F-actin might be organized to produce the required, concerted, force found in lamellipodial extensions of translocating cells.

      DISCUSSION

      Our conclusions rely upon use of quantitative measures of actin filament sliding within an in vitro motility assay system. Following our initial observation that filament sliding becomes strongly oriented when high surface densities of plain F-actin are present in this assay system, we defined a measure of orientedness based on the Kuiper statistic. This analytical tool enabled local domains of oriented motion to be identified so that the evolution of oriented patterns of movement could be followed in space and time. The fascinating aspect of the self-orienting system is that the length scale over which the patterns emerge are similar to the size of the leading lamella of a migrating mammalian cell. We found that oriented patterns of F-actin movement evolve with time and that the collision of differently oriented domains gives rise to new patterns of movement. This observation gives insight into how signals in one region of the leading lamellae might propagate across the cell via mechanical interaction between filaments so that cell migration might be steered toward or away from a chemical cue.
      At actin concentrations of above 20 μm we found that movement of rhodamine-phalloidin-labeled actin filaments became locally aligned and ordered domains formed in which movement was strongly oriented along one preferred axis. The degree and direction of alignment evolved with time and was clearly driven by the sliding motion generated by actomyosin interactions. A few tens of seconds after the addition of high background concentrations of plain F-actin; myosin driven movement of the filaments at the coverslip surface started to show increased speed and the direction of filament travel began to follow a straighter trajectory path. After a few minutes, movement of filaments within a typical field of view (100 × 100 μm) became strongly aligned along one preferred direction. The alignment direction was random with respect to microscope axis and once established, was maintained until Mg-ATP was depleted by actomyosin ATPase activity. In any particular experimental sample, discrete domains were observed with different alignment angles relative to each other. Domain alignments fluctuated on the second to minute time scale and variation in preferred angle with time depended upon the domain size. The degree of alignment of the population increased dramatically with actin concentration (and F-actin surface density) and the effect was maximal at concentrations of actin similar to those found in the leading lamella of the living cell (1 mg/ml = 25 μm actin).
      The behaviors reported here are in contrast to the sharp gel-sol transition of flow-aligned filaments measured in bulk solution (
      • Janmey P.A.
      • Hvidt S.
      • Käs J.
      • Lerche D.
      • Maggs A.
      • Sackmann E.
      • Schliwa M.
      • Stossel T.P.
      ). We found alignment depended upon filament length, filament surface density, and required the action of myosin molecular motors to drive the process. The surface alignment that we observe differs from solution flow-induced alignment in the following ways. 1) It is self-induced. 2) It is generated by actomyosin motility rather than thermal motion or bulk flow. 3) The aligned patterns extend over distances of only 100 μm and/or the direction of alignment is highly variable, so would average zero in a bulk measurement.
      We propose that the observed phenomena may mimic the dynamics of actin filaments at the rear region of the lamellipodium of migrating cells. The orientation of myosin molecules in our in vitro assays is random and yet the motion of actin was highly oriented and the preferred direction of travel could readily rotate with respect to the surface over a period of a minute. Myosin motors situated in the lamellipodium also probably have random orientation with respect to the axis of the cell yet they might produce highly directed bulk movement of actin due to lateral mechanical interactions between filaments. The density of myosin II in different cells varies widely: muscle myosin is present at 100 mg/ml in muscle cells, whereas non-muscle myosin IIa and IIb is present at about 0.2 mg/ml in platelets.
      J. R. Sellers, personal communication.
      We applied between 0.2 and 0.5 mg/ml of myosin (HMM or thick filaments), which gives an average density of myosin heads similar to that on a thick filament. So our assay conditions are not that dissimilar to the lamellipodium of a migrating cell in terms of the oligomeric state and concentration of myosin. Our observation that adjacent domains moving with different orientations interact and yield new patterns of motion over a period of a few seconds (e.g. Fig. 10) might be highly analogous to the mechanism underlying changes in direction of cell migration.
      Lamellipodia are packed with actin filaments and during cell migration the lamellipodial extension rapidly explores many orientations to identify the optimal one. In simplified cell motility assay systems (
      • Verkhovsky A.B.
      • Svitkina T.M.
      • Borisy G.G.
      ) it was discovered that application of small external forces applied using a micropipette were sufficient to guide the direction of translocating cytoplasts. This macroscopic phenomenon might be explained by the mechanism that we report here; applied external forces would perturb or add to the lateral forces between filaments that would then cause them to realign. A possible future avenue for study would be to try and forcibly align filaments within our assay either by local flow or local activation or inactivation of the myosin motors. Our present results imply that much of the dynamic behavior of lamellipodia might be controlled by alignment of sliding actin filaments in the myosin-rich region of the lamellipodium, located toward the rear region of the cell. This region has been proposed to be the “driving seat” where signals controlling cell migration are integrated (
      • Ridley A.J.
      • Schwartz M.A.
      • Burridge K.
      • Firtel R.A.
      • Ginsberg M.H.
      • Borisy G.
      • Parsons J.T.
      • Horwitz A.R.
      ). Cooperative, lateral mechanical interactions between actin filaments in this region of the cell might steer or bias the direction of lamellipodial protrusions further forward. Both the length and time scales of our observations seem completely consistent with this idea. Much is known about the dynamic nature of the actin meshwork at the leading edge of the cell (
      • Pollard T.D.
      ,
      • Pollard T.D.
      • Borisy G.G.
      ) and the current consensus is that actin polymerization in this region is the major force generator. Our proposal is that the direction of protrusion is steered by alignment of actin at the rear of the cell and that this process results from lateral mechanical interactions between filaments driven by myosin motors.
      Finally, it is noteworthy that the myosins (
      • Wylie S.R.
      • Chantler P.D.
      ,
      • Verkhovsky A.B.
      • Svitkina T.M.
      • Borisy G.G.
      ) thought to be involved in lamellipodial extrusion probably have a duty cycle ratio somewhere between that typical of processive motors like myosin V and the low duty ratio motors like muscle myosin II (
      • Wang F.
      • Kovacs M.
      • Hu A.
      • Limouze J.
      • Harvey E.V.
      • Sellers J.R.
      ,
      • Kovács M.
      • Wang F.
      • Hu A.
      • Zhang Y.
      • Sellers J.R.
      ). This means that actin filaments would be periodically released from the myosin during the ATPase cycle so that they could move laterally to explore new directions of travel in response to lateral forces exerted by near neighbor filaments.

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