TY - JOUR
AU - Watanabe, Naoki
AB - The retrograde actin flow, continuous centripetal movement of the cell peripheral actin networks, is widely observed in adherent cells. The retrograde flow is believed to facilitate cell migration when linked to cell adhesion molecules. In this review, we summarize our current knowledge regarding the functional relationship between the retrograde actin flow and focal adhesions (FAs). We also introduce our recent study in which single-molecule speckle (SiMS) microscopy dissected the complex interactions between FAs and the local actin flow. FAs do not simply impede the actin flow, but actively attract and remodel the local actin network. Our findings provide a new insight into the mechanisms for protrusion and traction force generation at the cell leading edge. Furthermore, we discuss possible roles of the actin flow-FA interaction based on the accumulated knowledge and our SiMS study. actin, cell migration, focal adhesion, retrograde flow, single-molecule speckle microscopy Cell migration is a dynamic, actin-based cellular process that is important for many phenomena in multicellular organisms, including development, wound healing, immunity and tumour metastasis. The motive power of cell migration is regulated by the actin cytoskeleton, which generates intracellular force by actin polymerization or interaction with myosin motor proteins. Although how cells use intracellular force in a spatiotemporally organized manner is a key question in cell migration research, it is challenging to monitor intracellular force directly in cells. Instead, monitoring motion of the objects that receive intracellular force may provide a clue to estimate dynamics of the force. The retrograde actin flow, continuous centripetal movement of the cell peripheral actin networks, has been observed in a wide variety of cultured cells, such as fibroblasts, epithelial cells, leukocytes, T lymphocytes and neurons (1–5). The physiological role of the retrograde flow largely remains unclear, although force driving the flow is considered to facilitate cell migration. During cell migration, a cell extends the lamellipodia, which is the sheet-like structure containing a highly dense, branched actin meshwork (6). The branched actin meshwork is generated by the Arp2/3 complex that binds to the side of a pre-existing filament and nucleates a new actin filament (7). In lamellipodia, actin filaments are mostly oriented with their barbed ends (plus ends) facing the cell edge (8, 9), and actin polymerization at the edge of lamellipodia generates force that pushes the membrane forward and pushes back the actin array inward to drive the retrograde flow (10, 11). During cell migration, the retrograde flow has been postulated to promote protrusion via linkage between the lamellipodial actin network and focal adhesions (FAs) in the clutch model (10). FAs are integrin-containing, multiprotein structures that form mechanical links between the extracellular substrate and actin stress fibres [reviewed in (12)]. FAs can also associate with lamellipodial actin networks via actin-binding FA components including vinculin, talin and α-actinin. In the clutch model, strengthening linkage between cell adhesion molecules and the lamellipodial actin network reduces the speed of the retrograde flow, whereas addition of new actin subunits continues at the barbed ends, leading to cell edge protrusions (10). The model also predicts that when the actin network moving with the retrograde flow links to FAs, tension is generated by flow motor(s) onto the substratum (10). Shootin-1, L1-CAM and N-cadherin have been proposed to function as a molecular clutch that link between the actin network and cell adhesion molecules in axon outgrowth and growth cone migration (13–15). However, it has been difficult to validate the original clutch model. To achieve this, it requires simultaneous measurements of the retrograde flow speed, the actin elongation rate at the cell edge and the rate of membrane protrusion with changing the strength of the linkage between FAs and the actin network. In addition, such biological processes are likely to be modulated by intracellular signaling, which is difficult to distinguish from the physical processes. Conversely, it has experimentally been suggested that when the substratum coated with the extracellular cell adhesion molecule is artificially contacted with the dorsal cell surface, linking the extracellular cell adhesion molecule to the cortical actin network increases tension between a cell and the substratum as well as membrane protrusion (16). Aplysia cell adhesion molecule (ApCAM) is a member of immunoglobulin-like cell adhesion molecules (Ig-CAMs), and its extracellular region mediates cell–cell adhesion by homophilic binding. Spectrin and ankyrin are candidates for actin-linkage molecules for Ig-CAMs via binding to both actin filaments and cytoplasmic domain of Ig-CAMs (17). When a silica bead coated with ApCAM is placed on the most distal dorsal surface of a growth cone of an Aplysia bag cell neuron and kept in one place using a glass needle, bending of the needle towards the cell centre, a decrease in the retrograde flow rate and the membrane protrusion anterior to the bead occur simultaneously (16). Bending of the needle indicates an increase in the tension on the beads. Thus the force that drives the retrograde flow is presumably transmitted to the extracellular beads accompanied by strengthening linkage between the actin network and the cell surface ApCAM. Tethering the actin network to the substrate via cell surface receptors may commonly generate tension on the substrate. Interestingly, such linkage between ApCAM beads and the actin network induces accumulation of actin and other molecules including Src and cortactin in the vicinity of the beads (16, 18, 19), suggesting additional intracellular signaling promotes protrusion. Nascent adhesions, which are the earliest integrin-containing structures detectable by light microscopy, are formed in lamellipodia (20, 21). Nascent adhesions contain major FA components including integrin, paxillin and α-actinin, and are roughly <0.25 µm in diameter (22). Most nascent adhesions disassemble rapidly, but a part of them grow into FAs by increasing their size and forming more stable association with the extracellular matrix (ECM) (23). As postulated in the clutch model, nascent adhesions and mature FAs are possible to link to the lamellipodial actin network moving with the retrograde flow (Fig. 1D). However, how these adhesions interact with the retrograde flow has been largely unknown. Morphology and distribution of mature FAs are diverse depending on the ECM proteins, the types of cells, stiffness of the substrate and so forth (Fig. 1). Under certain conditions, FAs are formed within lamellipodia. For instance, a part of FAs often exist in lamellipodia when XTC cells are cultured on a glass surface doubly coated with poly-l-lysine (PLL) and laminin (Fig. 1B). We have recently elucidated the relationship between the local retrograde flow velocity and FAs by direct observation of individual actin filaments using an improved fluorescence single-molecule speckle (SiMS) microscopy (24). Our study proposes a new idea that mature FAs attract the retrograde flow in lamellipodial region between the cell edge and the frontal edge of the FAs and actively remodel the local actin network. Fig. 1 Open in new tabDownload slide XTC cells display distinct actin and FA organization on different substrates. Cells express Lifeact-mCherry to visualize actin and EGFP-vinculin to mark FAs. Lamellipodia is the cell peripheral region that contains actin densely. (A) The cells form well-spread lamellipodia and few FAs on glass surfaces coated with 1 mg/ml PLL. (B) The cells form elongated FAs and spread lamellipodia on surfaces coated with 0.1 mg/ml PLL and 5µg/ml laminin. In these cells, edges of FAs frequently exist in lamellipodia. (C) The cells form narrow lamellipodia, FAs inside of lamellipodia, and well-developed actin stress fibres associated with FAs on surfaces coated with10 µg/ml each of PLL and fibronectin. Note that to prepare different substrates, the glass surfaces were incubated with the solution containing each substance at the indicated concentration, and then the excess amount of them was removed and washed away. (D) A schematic diagram of FAs and actin networks at leading edge of a cell. In this review, we discuss our current understanding of the physiological significance of the retrograde flow in cell migration. In particular, we highlight the relationship between the retrograde flow and FAs. We introduce imaging techniques to measure the flow velocities and our recent detailed analysis which revealed how FAs modify the local retrograde flow. Finally, we propose possible concepts that could explain how the interaction between the retrograde flow and FAs plays roles in organization of local cytoskeletal structures. How Do Cells Drive the Retrograde Flow in Lamellipodia? Adherent cells form diverse patterns of cytoskeletal and FA organization depending on the condition of culture substrates. Figure 1 shows examples of XTC cells showing distinct spatial arrangement of actin and FAs on different substrates. When XTC cells adhere to glass surfaces coated with 1 mg/ml PLL, they form well-spread lamellipodia and few FAs (Fig. 1A). Conversely, elongated FAs and spread lamellipodia are developed in XTC cells culture on surfaces coated with 0.1 mg/ml PLL and 5 μg/ml laminin (Fig. 1B). In these cells, edges of FAs frequently exist in lamellipodia (Fig. 1B). When XTC cells are cultured on 10 μg/ml each of PLL and fibronectin-coated coverslips, cells form narrow lamellipodia at the cell periphery, FAs inside of lamellipodia, and well-developed stress fibres associated with FAs (Fig. 1C). In these cells with distinct cytoskeletal organization, the retrograde actin flow ubiquitously occurs in lamellipodia. Based on the properties of the lamellipodial actin network with the free barbed-ends facing the cell edge and intensive polymerization at barbed ends, actin polymerization pushing against membrane tension is thought to partly drive the retrograde flow. In neuronal growth cones, application of cytochalasin B, which inhibits actin polymerization at barbed ends, causes receding of the actin network from the entire peripheral edge toward the cell body with forming a cell peripheral zone devoid of actin filaments (25). Thus, the retrograde flow is still driven without actin polymerization. A similar phenomenon is observed in sea urchin coelomocytes treated with cytochalasin D (26). In neural growth cones, the myosin II ATPase inhibitor blebbistatin decreases the retrograde flow in peripheral domain that corresponds to lamellipodia (27), but has little effect on the flow speeds in newly formed lamellipodia in T cells (28) and XTC cells on a PLL-coated surface (24). In neural growth cones, the actin bundles in the peripheral domain appear to be connected with the actin bundles containing myosin II in the central area of the growth cone (27), which could account for the sensitivity of its leading edge actin retrograde flow to myosin-II inhibition. Currently, the true flow motors that directly drive the lamellipodial actin flow remain to be identified. Interestingly, dendritic cells, a type of rapidly migrating leukocytes, seem to switch distinct mechanisms for effective migration in various extracellular environments. For instance, dendritic cells enables to migrate on ECM-coated adhesive substrate with forming a strong linkage between the actin network and the substrate, whereas they continue to migrate on non-adhesive substrate at the same speed without forming such linkage but with accelerated retrograde flow (3). In the latter case, when the cells migrate without adhesion, actin polymerization in the leading edge is accelerated to compensate, which presumably keeps membrane protrusion and migration velocity constant (3). Conversely, when Dictyostelium cells are detached from the substratum by changing the external solution, the actin filaments display accelerated retrograde flow, but the detached cells cannot advance pseudopods (29). Leukocytes are able to migrate flexibly through three-dimensional environments without tightly adhering to specific substrates (30). Therefore, such migration mode without clutch and rapid adaptation to environmental changes might be unique in leukocytes. Nevertheless, in other cell types, the retrograde flow and cell adhesion molecules may interact in a complex manner during cell migration. Direct Observation of the Modification of the Local Retrograde Flow by FAs The continuous centripetal flow of cell-associated material at the cell periphery is one of the most prominent phenomena in cultured cells, and has long been recognized with advances in microscopic techniques for live cell imaging. Abercrombie, who pioneered the study of cell migration [reviewed in (31)], first described the centripetal movement in chicken fibroblasts by observing particle transport on the cell surface using time-lapse phase-contrast microscopy (32). Wang (5) revealed the retrograde flow of the actin networks in lamellipodia by fluorescence recovery after photobleaching (FRAP) of fluorescently labeled actin. Later, fluorescent speckle microscopy (FSM) was developed with advances in high sensitivity cameras with cooled charge-coupled device detectors (33). FSM allows quantitative analysis to reveal the velocities of the retrograde flow with high resolution (4, 24, 34, 35). Linking the actin network to FAs in lamellipodia has been predicted to slow the speed of the retrograde flow locally. However, it has been difficult to detect the local modification of the actin flow by FAs especially when FAs are small. In addition, it is challenging to accurately measure the flow velocity of the actin population with short lifetime that moves only a short distance. Here, we introduce (i) several issues which one should keep in mind to accurately measure the actin flow velocity in lamellipodia, (ii) our improved speckle microscopy that overcomes those issues and (iii) our recent detailed analysis of the local retrograde actin flow near and above FAs with nanometre accuracy. Technical difficulties in measuring actin flow velocity Many studies have reported the actin flow velocity in lamellipodia using several microscopic techniques to reveal molecular dynamics such as FRAP (5), photoactivation of fluorescence (PAF; 36, 37), quantitative polarized light microscopy (Pol-Scope; 38) and FSM (4). However, it is difficult to accurately measure the flow velocities with the above techniques for following reasons. First, actin filament turnover in lamellipodia is rapid as nearly one-third of filaments have short lifetimes of less than 10 s (39). Such short-lived filaments move only short distances (<100–500 nm), thereby it requires a high spatial resolution to measure the flow velocity of these short-lived filaments. Second, if movements of actin filaments are heterogeneous, individual filaments must be tracked to define the flow. Approaches that monitor a mass of actin filaments, such as FRAP, PAF and Pol-Scope are not suited for measuring heterogeneous actin movements because of their limited spatial resolutions. FSM is an approach to monitor dynamics of cytoskeletal polymers in living cells with higher resolution than the above microscopic techniques. The principle of FSM is that incorporation of fluorescently labeled subunits at low concentrations provides fiduciary marks on polymers. FSM, which was originally developed by Waterman-Storer et al. (33) as a technique to measure microtubule movements and the actin flow, has been developed in two directions: one is fluorescence SiMS microscopy (24, 39) and the other is quantitative fluorescent speckle microscopy (qFSM; 40). In the original SiMS analysis, individual EGFP-actin molecules are imaged in cells expressing a very low level of EGFP-actin under the control of the defective cytomegalovirus (CMV) promotor (delCMV; 24, 39). The SiMS analysis enables to dissect complex actin dynamics, which is potentially overlooked by other approaches including FRAP, PAF and qFSM. However, this approach demands some experience to find cells containing fluorescent probes at optimal levels. Even with delCMV, the defective promoter, only a minor population of cells expresses a sufficiently low level of EGFP-actin. This technical difficulty might have limited usage of the method. qFSM analyses denser fluorophore labels than SiMS analysis (40). Resolvable fluorescent speckles containing multiple-fluorophore probes (2–10 fluorophores) provide quantitative information on appearance, disappearance and trajectories through automatic computational analysis (40). qFSM is a potent method with high statistical power because it extracts information from thousands of actin speckles within one image, yielding a heat map representation of degree of actin assembly/disassembly in subcellular areas. However, it could often be difficult to accurately measure the retrograde flow velocity by qFSM, because automated object tracking may misidentify individual speckles when they are densely packed. A remarkable example is that two studies in which researchers analysed the same FSM image datasets concluded differently (41, 42). One qFSM study proposed that actin filaments with two distinct retrograde flow rates coexist in lamellipodia of newt lung epithelial cells (41). This is referred to as the lamella hypothesis since the slower flow speed is equivalent to that in the area called lamella behind the back of lamellipodia. However, the other study failed to detect such slowly migrating fluorescent actin speckles in lamellipodia using several speckle tracking approaches (42). The contentious subject raised over the fidelity of qFSM has been still an unsettled issue (42, 43). Tracking the retrograde flow at nanometre scales by new SiMS microscopy We recently developed a new, user-friendly method, which achieves the SiMS imaging with the highest spatiotemporal resolution (24). The outline of the new method is shown in Fig. 2A. We use an actin probe labeled with a fluorescent DyLight dye (Thermo Fisher Scientific Inc.) on lysine side chains. To deliver DyLight (DL)-actin into cells, we use electroporation, which enables incorporation of DL-actin into the cytoplasm at a low density suitable for SiMS microscopy (Fig. 2B and C; 24). With our electroporation method, almost 100% of cells are labeled with DL-actin molecules at a similar density. In cells, DL-actin efficiently incorporates into cellular actin networks, and especially, DL549- and DL550-actin show greatly improved photostability and brightness. We took advantage of these properties of DL-actin to increase the accuracy of displacement measurements of actin speckles by devising two strategies. First, the unattenuated 100-W mercury-arc lamp is used for illumination. Mercury-arc has a strong emission peak at 546 nm, which enables monitoring of DL-actin SiMS with a high signal-to-noise ratio even with short exposure time. Typically, we acquire images at 100 ms intervals (fast tracking imaging) for the short duration of 10 s (Fig. 2C). Note that special care must be taken to minimize the photodamage by restricting the illuminated area and duration of the fast tracking imaging in live cells. A decrease in the retrograde flow shows a visible sign of the photodamage (24). With our microscope system, we restrict duration for the fast tracking imaging within 10 s. Second, subpixel localization of the SiMS centroid is determined using the two-dimensional Gaussian fit model of Speckle TrackerJ software (44). Under this condition, the centroid of DL-actin SiMS can be resolved with a localization error of 8–8.5 nm (24). Thus in vivo nanometre-scale robust velocity measurement of SiMS whose displacement is only 100–150 nm (Fig. 2D), i.e. within the diffraction limit of light microscope, is achieved. Fig. 2 Open in new tabDownload slide New SiMS microscopy using DyLight-actin (DL-actin) is user-friendly and enables in vivo nanometre-scale displacement analysis. (A) Outline of the new SiMS method. (B) Images of DL549-actin speckles in lamellipodia of XTC cells. Time-lapse images paneled at 10 s intervals in the area (square) are shown on the right. Bar, 10 µm. (C) An image of DL549-actin with fast tracking method. The image was acquired with a 100 ms exposure time and a full 100-W mercury excitation. Bar, 5 µm. (D) Displacement plot of the central position of a short-lived DL549-actin speckle in lamellipodia in the series of fast tracking images. The velocity was calculated from a linear fit. Modified from Yamashiro et al. 2014 (24). Using the new SiMS method, we, for the first time, succeeded in the measurement of the flow rates of actin speckles including short-lived species (lifetime <10 s), which move only a few hundred nanometre distance (24). In lamellipodia of XTC cells shortly after spreading on PLL-coated coverslips, all DL-actin SiMS including short-lived species flowed at uniform speeds, suggesting that lamellipodial actin filaments form a tightly connected network regardless of individual filament lifetimes. Thus, the lamella hypothesis (41) does not apply in lamellipodia of XTC cells. New SiMS microscopy dissects local retrograde flow near FAs Using the improved SiMS microscopy, we analysed how FAs influence local retrograde flow in lamellipodia in detail (24). As shown in Fig. 1B, when XTC cells are spread on a substrate coated with PLL and laminin, nascent adhesions and mature FAs are formed in flat lamellipodia, and a part of FAs frequently exist in lamellipodia of the cells. Thus, we used the cells under the condition to study the relationship between the retrograde flow in lamellipodia and FAs. Nascent adhesions are the first observable adhesive structures, which are small and highly dynamic (12). In SiMS analysis with DL-actin, the speeds of actin speckles flowing over nascent adhesions are similar to those of speckles that flowed near the nascent adhesions (Fig. 3A). In our study, the speed of DL-actin speckles flowing over nascent adhesions was 97.3 ± 0.06% of that of speckles flowing near but outside of the adhesions (n = 15 cases of comparison, four cells; 24). These results suggest that nascent adhesions barely link to the lamellipodial actin networks moving along the retrograde flow (24). Fig. 3 Open in new tabDownload slide Effects of nascent adhesions and mature FAs on the local retrograde flow at the cell periphery. (A) A speed and trajectory map of the DL549-actin speckles in lamellipodia containing nascent adhesions. The inset indicates images of EGFP-vinculin before (green) and after (red) acquisition of the DL549-actin movie. Lines indicating the trajectories and average speeds of DL549-actin speckles observed within a 100-s-time window are shown in an image of EGFP-vinculin. Nascent adhesions have little effect on local retrograde flow at the cell periphery. (B) An average speed and trajectory map of DL549-actin speckles in lamellipodia containing a mature FA. A white dotted line outlines the FA. Lines indicating the trajectories and average speeds of DL549-actin speckles as in A. (C) A local speed and trajectory map of representative DL549-actin speckles in the boxed region in (B). Circles indicate locations of speckles in each frame. Local actin flow speeds were measured with a 5-s-time window (five frames), and the circles corresponding to the intermediate time point (the third frame) were coloured according to speed. Green arrows indicate the direction of the retrograde flow. Asterisks indicate the speckles slowed down at the frontal edge of FA. Arrowheads indicate the speckles migrating beside FA. An open arrowhead indicates a speckle migrating beside FA and changing its direction. Modified from Yamashiro et al. 2014 (24). In contrast, mature FAs locally modify the retrograde flow in several ways. (i) In the centre of FAs, actin speckles move slower than those flowing in other parts of lamellipodia (Fig. 3B, the region surrounded by a white break line). (ii) In the cell peripheral region between the cell edge and the frontal edge of FAs, the retrograde flow biases towards FAs (Fig. 3B). (iii) Also, in the cell peripheral region between the cell edge and the frontal edge of FAs, the actin speckles move at faster speeds than those elsewhere in lamellipodia (Fig. 3B). Detailed local flow speed analyses show that the speckles flowing into the FA area often gradually slow down as the speckles approach to FAs (Fig. 3C, the speckle trajectories with asterisks). Based on (i)–(iii), mature FAs appear to accelerate and gather the retrograde flow in front of FAs and locally drag the actin flow at the frontal edge of the FAs. (iv) The speckles migrating beside FAs move at a constant speed (Fig. 3C, the speckle trajectories with filled and open arrowheads), and sometimes change their direction without changing the speed (Fig. 3C, the speckle trajectory with open arrowhead), suggesting that they dodge the FAs. Presumably, the mature FAs physically impede the actin flow. (v) The speckles migrating on the lateral edge of FAs occasionally move away from the side of FAs with changing their speeds irregularly (Fig. 3C, the speckle trajectories coloured with light blue). The actin network in the lateral edge of FAs might be partly connected to and pulled out by the actin networks outside of the FAs. We reproducibly observe the above features of local retrograde flow near mature FAs in XTC cells (24). The observed modification of the actin flow velocity by FAs must involve transformation of the actin network near and within FAs. The fast-moving actin networks flowing towards FAs are transformed into slow-moving networks. This transformation is presumably caused by engagement between the actin network and FAs, or local changes in force to drive the actin flow. Conversely, part of the actin network flowing towards the side of FAs seems to dodge the FAs and flow at an unaltered speed. The rapid turnover of lamellipodial actin networks might facilitate the flexible transformation of the network locally to adapt for obstruction by FAs. Possible Roles of Gathering and Collecting Action of FAs on the Retrograde Flow Based on the above observations, we conclude that mature FAs attract the flow in front and actively remodel the local actin network. Although the mechanism for modification of the retrograde flow by FAs is still unclear, we propose three possible roles of the actin flow-FA interaction based on our SiMS study (24) and the knowledge from previous studies. The retrograde flow may recruit components of FAs The most possible role of the retrograde flow is to recruit FA components to FAs for maintenance (Fig. 4, Model-I). In mature FAs, a constant turnover of FA components, such as vinculin and paxillin, on timescales of seconds has been demonstrated using FRAP (45). The retrograde flow streaming into FAs could effectively supply FA components including actin-binding proteins, which bind directly or indirectly to the lamellipodial actin network. In addition, the retrograde flow may remodel ECM via binding to integrins in front of FAs. The ECM is a dynamic structure undergoing remodeling processes, which are involved in morphogenesis and diseases including tissue fibrosis and cancer invasion [reviewed in (46,) and (47)]. Indeed, live imaging with fluorescent fibronectin during cell migration in Xenopus embryonic tissue explants shows that rearrangement of fibronectin fibrils occurs as the cells migrate (48). These possibilities should be addressed by observing behaviours of the representative FA components and ECM in front of FAs. The SiMS microscopy would be a suitable approach to address such possibilities. Fig. 4 Open in new tabDownload slide Possible roles of gathering and collecting action of mature FAs on the retrograde flow in front of FAs. In the Model-I, the retrograde flow recruits FA components which bind directly or indirectly to the lamellipodial actin network to FAs. In the Model-II, FAs increases the traction force and extend the region at the frontal edge of FAs by modification of the retrograde flow. In the Model-III, the retrograde flow remodels the lamellipodial actin network with the barbed-ends facing the cell edge to anti-parallel actin bundle in stress fibres. When part of an actin filament near the barbed end links to a FA, the pointed end part of the filament flows faster than the other part of the filament, thereby turns and the filament align along the orientation of the actin flow (left). Conversely, when part of an actin filament near the pointed end links to a FA, the barbed end part of the filament rotates so that the pointed end of the filament is directed toward the cellular edge (right). In this way lamellipodial actin filaments might be reorganized to anti-parallel bundles. In the Model-IV, mechanical stress caused by collision of the actin network accumulates in the forepart of FAs. In this model, such mechanical stress promotes disassembly of actin filaments locally, whereas part of actin filaments oriented along the flow direction are stabilized by tension in FAs. The retrograde flow modified by FAs may increase the traction force at the frontal edge of FAs Deceleration of the retrograde flow in the region where the flow runs into FAs may result from the linkage between the actin network and mature FAs. In this case, traction force on the substratum can be exerted by the retrograde flow-FA interaction. Dynamic traction force within individual mature FAs on the substratum has been analysed using high-resolution traction force microscopy (49, 50). An inverse relationship between the retrograde flow speed and traction force on the substratum in lamellipodia and lamella-containing FAs have been demonstrated using combination of FSM and traction force microscopy (49). Furthermore, the traction force decreases but remains in the blebbistatin-treated cells in which the retrograde flow remains in lamellipodia, whereas the actomyosin contractility in stress fibres is inhibited (49). These observations suggest that the traction force is generated when the actin network moving with the retrograde flow links to FAs. Several modeling studies presumed a ‘stick-slip’ motion in which the actin network-FA interaction occurs intermittently (51, 52). It is also predicted that rubbing nascent adhesions by the actin network moving with the retrograde flow reinforces the link between integrins and actin filaments through modifying tension-sensitive molecules include talin, p130Cas, fibronectin and integrin (12, 53), thereby promotes growth of nascent adhesions into FAs. These models assume that FAs simply produce friction to the retrograde actin flow. Conversely, it has not yet been demonstrated rigorously whether the retrograde flow promotes growth of nascent adhesions in cells. Inhibition of actin polymerization by cytochalasin D diminishes both the retrograde flow and nascent adhesions (53). However, since cytochalasin D diminish the structure of the actin network at the lamellipodium tip (25, 26), the effect on nascent adhesions might be caused by the collapse of the surrounding actin structure. In our SiMS study, mature FAs appear to actively remodel the local actin network to accelerate and gather the retrograde flow in front of FAs (24). This act of the FAs would lead to increase in traction force and extend the region in where traction force is exerted at the frontal edge of FAs (Fig. 4, Model-II). If a new technique to measure traction force at growing adhesion structures with sufficient resolution is developed, we might be able to evaluate the correlation between change in the flow speeds, traction force and accumulation of FA components during assembly of FAs. Modification of the retrograde flow by FAs may remodel lamellipodial actin filaments to stress fibres As described in Introduction, lamellipodia contain a dense, branched meshwork of actin filaments with their barbed ends directed towards the leading edge of the cell (6). Part of lamellipodial actin filaments might be reorganized into FA-associated actin stress fibres, which are composed of bundled actin filaments associated with myosin II and actin cross-linking proteins. Actin stress fibres contain anti-parallel filaments that contribute to myosin II-dependent contractility. We propose a model in which the FA-modulated local retrograde flow promotes the redirection of lamellipodial filaments into anti-parallel filament bundles. In our SiMS study, actin filaments above mature FA moved at slow speeds probably due to the friction between actin filaments and FAs, whereas actin filaments at the side of FAs flows at fast speeds (Fig. 3B; 24). As shown in Fig. 4 (Model-III), when part of an actin filament near the barbed end links to a FA, the pointed end part of the filament turns and the filament align along the orientation of the retrograde flow (Fig. 4, Model-III, left). Conversely, when part of an actin filament near the pointed end links to a FA, the barbed end part of the filament might rotate so that the pointed end of the filament is directed towards the cell edge (Fig. 4, Model-III, right). These actin filaments oriented in an anti-parallel manner may be cross-linked to form actin bundles in stress fibres. Mechanical stress caused by collision of the actin network moving at a fast flow speed in front of FAs and that moving at a slow speed over FAs could accumulate at the frontal edge of FAs (Fig. 4, Model-IV). Such mechanical stress may promote disassembly of actin filaments locally. Currently, there is neither biochemical nor cell biological evidence whether mechanical stress promotes disassembly in the Arp2/3 complex-mediated actin meshwork. Nevertheless, we recently show that mechanical stress applied by microneedle manipulation causes a rapid actin monomer increase in XTC cells (54). Mechanical stress also increases the level of actin-associated AIP1, which enhance actin-depolymerizing factor (ADF)/cofilin-mediated actin disassembly (54). In addition, a previous qFSM study reported that actin–myosin II contraction enhances actin disassembly in keratocytes (55). Conversely, part of actin filaments, especially these oriented along the retrograde flow direction in the forepart of FAs, can be stabilized by tension. A biochemical study suggests that tension along an actin filament reduces the binding of ADF/cofilin, thereby preventing the filament from being severed by those proteins (56). In this way, actin filaments flowing into FAs might survive and form actin bundles. It is intriguing to see how actin structures are remodeled accompanied by the modulation of the local retrograde flow. The combination of the SiMS microscopy and the EM tomography would serve to address the model (Fig. 4, Model-III). The recent electron tomography is capable of distinguishing polarity of actin filaments in cells (8), which should help examine the orientation of actin filaments near/in FAs. Conclusion We have summarized current knowledge regarding the relationship between the retrograde actin flow and FAs. FAs do not simply impede the local retrograde flow, but actively attract and remodel the local actin network. Our findings provide a new insight into the mechanisms for protrusion and traction force generation at the cell leading edge. Further studies using our new SiMS microscopy and combination with other microscopic techniques will elucidate how cells generate intracellular force and transmit the force to the extracellular matrix during migrating. Acknowledgements This work was supported by NEXT program grant LS013 from the Cabinet Office, Government of Japan (N.W.), a grant from Takeda Science Foundation (N.W.) and a Grant-in-Aid for Scientific Research on Innovative Areas Grant Number 00624347 from the Ministry of Education, Science, Sports and Culture of Japan (S.Y.). Conflict of Interest None declared. 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TI - A new link between the retrograde actin flow and focal adhesions
JF - The Journal of Biochemistry
DO - 10.1093/jb/mvu053
DA - 2014-11-01
UR - https://www.deepdyve.com/lp/oxford-university-press/a-new-link-between-the-retrograde-actin-flow-and-focal-adhesions-OB0NObimKE
SP - 239
EP - 248
VL - 156
IS - 5
DP - DeepDyve
ER -