Cells use a large repertoire of proteins to remodel the actin cytoskeleton. Depending on the proteins involved, F-actin is organized in specialized protrusions such as lamellipodia or filopodia, which serve diverse functions in cell migration and sensing. Although factors responsible for directed filament assembly in filopodia have been extensively characterized, the mechanisms of filament disassembly in these structures are mostly unknown. We investigated how the actin-depolymerizing factor cofilin-1 affects the dynamics of fascincrosslinked actin filaments in vitro and in live cells. By multicolor total internal reflection fluorescence microscopy and fluorimetric assays, we found that cofilin-mediated severing is enhanced in fascin-crosslinked bundles compared with isolated filaments, and that fascin and cofilin act synergistically in filament severing. Immunolabeling experiments demonstrated for the first time that besides its known localization in lamellipodia and membrane ruffles, endogenous cofilin can also accumulate in the tips and shafts of filopodia. Live-cell imaging of fluorescently tagged proteins revealed that cofilin is specifically targeted to filopodia upon stalling of protrusion and during their retraction. Subsequent electron tomography established filopodial actin filament and/or bundle fragmentation to precisely correlate with cofilin accumulation. These results identify a new mechanism of filopodium disassembly involving both fascin and cofilin.

Introduction

The dynamic rearrangement of actin filaments is fundamental for cell motility, and cells use a plethora of proteins to precisely regulate and coordinate the polymerization and depolymerization of actin filaments (Pollard and Cooper, 2009; Bugyi and Carlier, 2010). Although much has been learnt about the cellular players that mediate actin filament nucleation and elongation in protrusive structures such as lamellipodia and filopodia (Chesarone and Goode, 2009; Insall and Machesky, 2009), little is known about the mechanisms of filament disassembly.

One of the key players in filament depolymerization and actin monomer recycling is the actin depolymerization factor cofilin (Bernstein and Bamburg, 2010). Cofilin preferentially binds to ADP–F-actin and mediates severing and depolymerization by altering the mechanical properties of the filament: structural and biochemical analyses revealed that cofilin binding changes the subunit tilt and increases the helical twist of the filament (McGough et al., 1997; Galkin et al., 2001), thereby weakening lateral contacts between actin monomers (McGough and Chiu, 1999; Bobkov et al., 2004; Paavilainen et al., 2008) and considerably increasing the elasticity of the filament (Prochniewicz et al., 2005; McCullough et al., 2008).

In cells, cofilin is associated with the entire lamellipodium (Lai et al., 2008) and in one model promotes actin filament treadmilling in the lamellipodium and in Listeria comet tails by mediating filament disassembly of aged actin filaments (Loisel et al., 1999; Iwasa and Mullins, 2007; Kiuchi et al., 2007; Lai et al., 2008). Consistently, cofilin is an essential component of reconstituted motility medium (Loisel et al., 1999). However, in contrast to its established role in the turnover of lamellipodial actin filaments, a potential function in the dynamics of filopodia has so far been elusive. Filopodia are spiky, actin-rich protrusions that frequently emerge from the leading edge of the cell, and serve as sensory organelles, e.g. in axon guidance or endothelial zippering (Dent et al., 2007). Importantly, the fast growing, barbed ends of the filaments packed together in a filopodium point towards the membrane, producing enough force to push the membrane outwards. There is much debate about the mechanisms of filopodia initiation, but it is commonly believed that protrusion and retraction of these structures are effected by modulating the insertional assembly of actin monomers at their tips, and by inducing filament disassembly, potentially mediated by cofilin, at their base (Gehler et al., 2004; Mattila and Lappalainen, 2008; Faix et al., 2009; Hotulainen and Hoogenraad, 2010). In addition, actin filaments in filopodia are compacted into dense bundles by specialized proteins such as fascin, which localizes along their shafts (Otto et al., 1979; DeRosier and Edds, 1980), enhancing their rigidity and thus promoting their protrusion from the cell body (Vignjevic et al., 2006; Lieleg et al., 2007). Notably, fascin appears to undergo frequent cycles of association and dissociation with actin bundles both in vivo and in vitro instead of stably binding to filaments, which might be an important feature in regulating the dynamic growth and shrinkage of filopodia (Vignjevic et al., 2006; Aratyn et al., 2007). Moreover, fascin was shown to induce overtwisting of actin filaments within bundles, resulting in bundles with a hexagonal geometry consisting of ~20 actin filaments at maximum (Claessens et al., 2008).

We investigated the molecular details of the interplay of fascin and cofilin during actin assembly and disassembly with a combined approach, using multi-color total internal reflection fluorescence (TIRF) microscopy with purified proteins, fluorimetric assays and correlated live-cell imaging and electron microscopy (EM) tomography. We found that cofilin-mediated filament severing is enhanced in fascin-bundled actin filaments, and that cofilin can efficiently disassemble fascin-crosslinked actin bundles in vitro. Furthermore, we show that cofilin has an unexpected role in disassembly of filopodia, as it massively accumulates in tip and shaft regions of these protrusions, resulting in their rapid retraction and disassembly.

Results

Fascin mediates rapid crosslinking of actin filaments into parallel bundles

Initially, we employed in vitro TIRF microscopy of polymerizing, freely diffusing actin filaments to visualize the dynamic process of fascin-mediated actin bundle formation. This setup made it possible to discriminate between the fast growing barbed end and the slowly growing pointed end of the filaments and therefore to unambiguously determine the polarity of the formed bundles, as well as to determine the number of filaments per bundle and to analyze the kinetics of bundle formation. Single actin filaments that grew in the presence of fascin frequently fused with other filaments, resulting in a network of actin filament bundles (Fig. 1A; supplementary material Movie 1). The elongation rate of actin filaments was not altered by fascin and was ~11 subunits/second for TIRF experiments using 1.3 μM actin (30% Alexa-Fluor-488 labeled). The fluorescence intensities of the actin bundles correlated directly with the number of actin filaments within the bundle, which could readily be determined by following the time-resolved fusion of single actin filaments with each other and with other bundles (Fig. 1B). The number of filaments per bundle was largely independent of the fascin concentration used in our assay, as a concentration range from 10 nM to 10 μM fascin produced bundles consisting of approximately three to six actin filaments at steady state, with a maximal average number of filaments per bundle at 500 nM fascin (Fig. 1C; supplementary material Fig. S1). Although fascin is known to restrict bundle thickness to ~20 filaments (Claessens et al., 2008), the small effect of fascin concentration on bundle thickness observed here is probably caused by the low total amount of actin filaments in these assays, limiting the number of actin filaments available for bundle formation. Nevertheless, the total amount of bundled actin increased with fascin concentration, but reached a maximum at 50–100 nM fascin in the reaction mixture. This was true for both TIRF microscopy and low-speed sedimentation assays of actin bundles followed by densitometry after SDS-PAGE (supplementary material Fig. S1).

The analyses of the dynamics of fascin-mediated bundle formation showed some striking effects: first, the bundles formed became very rigid after fusion of two to three filaments, resulting in the appearance of numerous ‘kinks’ (supplementary material Movie 1), and the force produced by fascin-mediated stiffening of the bundle was apparently high enough to occasionally break existing bundles (supplementary material Movie 2). Second, analysis of the directionality of single growing filaments within the bundles revealed that all bundles formed by fascin were exclusively composed of parallel actin filaments (Fig. 1D; supplementary material Movie 2). Third, single filaments were rapidly fused to the actin bundle after coming in close proximity, a process we will refer to herein as ‘zippering’ (Fig. 1E; supplementary material Movie 3). This process was remarkably rapid, exceeding velocities of 700 subunits per second (Fig. 1E), which corresponds to the fusion of more than 100 μm actin filament/minute. Thus, the rate of fascin-mediated bundle formation is considerably faster than both actin filament elongation rates in vitro and protrusion rates of 2–30 μm/minute of filopodia and microspikes seen in different cell types (Sheetz et al., 1992; Costantino et al., 2008; Geraldo and Gordon-Weeks, 2009), and thus likely not rate-limiting during filopodia formation.

Cofilin efficiently severs actin filaments in fascin-crosslinked bundles

A previous study showed that spontaneously formed actin bundles were more resistant to cofilin-mediated severing than single filaments in vitro (Michelot et al., 2007). However, the effects of filopodial bundling proteins such as fascin on cofilin-mediated filament severing were not investigated. Therefore, we analyzed the dynamics of actin networks formed in the presence of both fascin and cofilin. By using single filament TIRF microscopy, we show that cofilin alone efficiently severs growing actin filaments with a robust activity at 300 nM in our assay (Fig. 2A; supplementary material Movie 4), whereas concentrations below 200 nM did not induce substantial filament severing (data not shown). Monitoring cofilin-mediated severing of polymerizing filaments instead of filaments at steady state has the advantage of allowing the unambiguous identification of severing events resulting from growth of newly formed barbed ends. We next directly visualized cofilin-binding to growing actin filaments using two-color TIRF microscopy with Alexa-Fluor-633-labeled actin and EGFP-tagged cofilin: the latter having been previously shown to have similar activities in vitro to the untagged protein (Lai et al., 2008). Comparison of the severing efficacies revealed that both untagged and EGFP-tagged cofilin are equally effective (Fig. 2B). In order to further confirm the functionality of EGFP-tagged cofilin, we performed fluorescence titrations with pyrene-labeled F- and G-actin and found that both cofilin variants exhibited comparable affinities for both actin species (supplementary material Fig. S2). Thus, EGFP-tagged cofilin is suitable to directly visualize actin-filament binding and severing. Two-color TIRF microscopy revealed that decoration of actin filaments with EGFP–cofilin was barely detectable in a region up to ~2.8 μm behind the growing barbed end, whereas it markedly decorated further proximal, aged actin filaments (Fig. 2B; supplementary material Fig. S3A and Movie 5), consistent with recent work using an Alexa-Fluor-488-labeled yeast cofilin mutant (Suarez et al., 2011). Interestingly, filament decoration with EGFP–cofilin occurred in a discontinuous fashion, indicating that side binding of cofilin to F-actin is a cooperative process, as previously proposed (supplementary material Fig. S3B) (Cao et al., 2006; De La Cruz, 2005; Ressad et al., 1998). Moreover, the simultaneous imaging of growing filaments and EGFP–cofilin allowed us to analyze the time-course of cofilin binding. Because actin filaments grew at 11 subunits/second in our assay, we calculated that EGFP–cofilin preferably binds actin filaments older than ~2 minutes, which is in line with a slow rate of phosphate release from ADP+Pi actin in the filament (Carlier et al., 1988), as well as with the selectivity of cofilin for binding to ADP–F-actin (Maciver et al., 1991; Michelot et al., 2007). However, sporadic EGFP signals could also be detected within the distal 2.8 μm of growing filaments (supplementary material Fig. S3). By analyzing the fragment lengths of cofilin- and EGFP–cofilin-severed filaments, we found that severing was most effective in a region of ~4.5–5 μm behind the growing barbed end. Importantly, the lengths of severed barbed-end fragments in the presence of 300 nM EGFP-tagged and untagged cofilin were virtually identical, affirming that the EGFP tag does not noticeably interfere with F-actin binding or concomitant severing. The majority of severed fragments were 3–10 μm in length, and several EGFP–cofilin decorated filaments where not severed at all during the experiments (Fig. 2D; supplementary material Movie 5). These findings are in good agreement with recent observations (Suarez et al., 2011) and corroborate a model predicting that cofilin severing activity is maximal when only parts of the filament are bound by cofilin, whereas a complete decoration results in decreased filament severing (De La Cruz and Sept, 2010).

Fig. 1.

Dynamics of fascin-mediated actin filament bundling. (A) Polymerization of 1.3 μM actin (30% Alexa-Fluor-488 labeled) in the presence of 500 nM fascin in TIRF buffer. Time is indicated in seconds in the top right of each panel. (B) The fluorescence intensity of the bundle directly correlated with the number of filaments. (C) The number of filaments per bundle formed in the presence of different fascin concentrations. For each condition, at least 30 bundles were analyzed using fluorescence intensity measurements. Boxes indicate 25th percentile, median and 75th percentile of all values; error bars indicate 10th and 90th percentile; minimal and maximal fluorescence intensities are also indicated. The number of filaments per bundle at steady state was only slightly increased at higher fascin concentrations. (D) Time-lapse micrographs of the formation of fascin-crosslinked F-actin bundles. Fascin apparently enhanced the stiffness of the bundles, resulting in kinks and breaks (arrows). All actin filaments within the bundles were oriented in the same direction. Time is indicated in seconds. (E) Time-lapse micrographs of the ‘zippering’ of actin filaments (arrows). The length of the zippered region was measured over time, yielding zippering velocities of ~700 subunits/second. Time is indicated in seconds. Scale bars: 5 μm (A,B,D,E).

Fig. 1.

Dynamics of fascin-mediated actin filament bundling. (A) Polymerization of 1.3 μM actin (30% Alexa-Fluor-488 labeled) in the presence of 500 nM fascin in TIRF buffer. Time is indicated in seconds in the top right of each panel. (B) The fluorescence intensity of the bundle directly correlated with the number of filaments. (C) The number of filaments per bundle formed in the presence of different fascin concentrations. For each condition, at least 30 bundles were analyzed using fluorescence intensity measurements. Boxes indicate 25th percentile, median and 75th percentile of all values; error bars indicate 10th and 90th percentile; minimal and maximal fluorescence intensities are also indicated. The number of filaments per bundle at steady state was only slightly increased at higher fascin concentrations. (D) Time-lapse micrographs of the formation of fascin-crosslinked F-actin bundles. Fascin apparently enhanced the stiffness of the bundles, resulting in kinks and breaks (arrows). All actin filaments within the bundles were oriented in the same direction. Time is indicated in seconds. (E) Time-lapse micrographs of the ‘zippering’ of actin filaments (arrows). The length of the zippered region was measured over time, yielding zippering velocities of ~700 subunits/second. Time is indicated in seconds. Scale bars: 5 μm (A,B,D,E).

To test whether the selectivity of cofilin for aged actin filaments persisted in polymerizing, fascin-crosslinked bundles and whether and how these bundles can be efficiently severed by cofilin, we first initiated the polymerization of actin in the presence of fascin and allowed the reaction to proceed until several growing bundles were formed and their barbed ends identified. Subsequently, cofilin was added to the reaction mixture, resulting in rapid disassembly of aged bundles, whereas short, bundled barbed-end fragments persisted (Fig. 2C,D; supplementary material Movies 6 and 7). In addition, the majority of bundled barbed-end fragments formed immediately after cofilin addition was only ~2 μm in length, and hence significantly shorter on average than the barbed end fragments produced by stochastic cofilin severing of individual actin filaments in the absence of fascin (Fig. 2D). The lengths of the severed barbed-end fragments in the presence of fascin correlated reasonably well with the lengths of filament fragments not decorated by cofilin in the absence of fascin (Fig. 2B,D), suggesting that filament severing is highly efficient in fascincrosslinked bundles and that it occurs immediately after cofilin binding. Most importantly, the generated bundled barbed ends continuously increased in fluorescence intensity, indicative of severing and continuous growth of barbed ends formed in these conditions (Fig. 2C,E; supplementary material Movies 6 and 7). However, the fact that the polarity of these bundles still persisted suggests that fascin crosslinking is not directly affected by cofilin. Notably, the bundles did not show a uniform increase in fluorescence, but displayed a varying but characteristic fluorescence distribution along their length: although the first 1–2 μm at the barbed end of the growing bundle contained only few filaments, upon further polymerization, the number of filaments increased substantially in a region 2–8 μm behind the bundle tips. Furthermore, the fluorescence intensity in the proximal ends of the bundles slightly decreased again with increasing amounts of bound cofilin (supplementary material Movie 6). Colocalization of EGFP–cofilin colocalization of EGFP–cofilin and Alexa-Fluor-633–actin confirmed that cofilin preferentially binds to aged F-actin within the bundle, as is the case for individual filaments (Fig. 2B), because it starts to accumulate in a region ~2–3 μm behind the growing bundle tip (supplementary material Fig. S4 and Movie 8). Most notably, the maximal fluorescence of actin and cofilin within fascincrosslinked bundles did not correlate directly, but instead the peak of cofilin fluorescence lagged behind that of actin by ~3 μm (supplementary material Fig. S4 and Movie 8). Thus, the data indicate that low amounts of cofilin rapidly sever actin filaments upon initial binding, resulting in formation of new, growing barbed ends within the bundle in these conditions, and that increasing amounts of cofilin in the rear of the bundle eventually promote its disassembly (supplementary material Movies 7 and 8). Subsequent analyses revealed that the prominent bundle thickening depended on the cofilin concentration and that robust thickening of fascin-crosslinked bundles even occurred at concentrations as low as 50–200 nM, which is not sufficient to efficiently sever single actin filaments in the absence of fascin in our assays (Fig. 2F; supplementary material Fig. S5A; and data not shown). Addition of 500 nM cofilin enhanced the calculated maximal number of filaments per bundle from between five and ten filaments to more than 100 as estimated by fluorescence intensity measurements (Fig. 2F). We subsequently analyzed the time course of cofilin binding to pyrene-labeled F-actin crosslinked by different amounts of fascin to analyze whether cofilin binding to crosslinked filaments might be impaired. Binding of cofilin to F-actin causes a decrease in pyrene fluorescence, providing a means of analysing the binding kinetics (Blanchoin and Pollard, 1999). The rate of cofilin-binding indeed decreased with increasing amounts of fascin in the bundling reaction (Fig. 2G). However, the overall change in fluorescence was only modestly lowered at increasing fascin concentrations compared with non-crosslinked filaments, suggesting that the slower binding kinetics result from impaired diffusion into the actin bundles or lower association rates of cofilin to fascin-decorated filaments rather than direct competition of fascin and cofilin for F-actin binding sites. Consistently, low-speed sedimentation assays revealed that the amount of fascin in F-actin bundles is not reduced by cofilin (supplementary material Fig. S5B). Taken together, these results suggest that cofilin severing activity is greatly enhanced in fascin-crosslinked bundles despite lower binding kinetics of cofilin when compared with individual filaments in the absence of fascin.

Fig. 2.

Cofilin-mediated severing of individual filaments and fascincrosslinked bundles. (A) 1.3 μM actin (30% Alexa-Fluor-488 labeled) was polymerized in the presence of 300 nM cofilin in TIRF buffer. The time-lapse micrographs show frequent severing of actin filaments (arrowheads). Time is indicated in seconds. (B) Left: direct visualization of EGFP–cofilin (green) binding to polymerizing Alexa-Fluor-633-labeled actin filaments (30% labeled; red). EGFP–cofilin preferentially bound to and decorated aged actin filaments, resulting in severing (arrowheads), whereas the barbed-end region was neither markedly decorated with EGFP–cofilin nor severed (asterisks). Time is indicated in seconds. Scale bar: 10 μm (A,B). Right: quantification of severing by cofilin and EGFP–cofilin. (C) Cofilin-mediated severing of fascin-bundled filaments. 1.3 μM actin (30% Alexa-Fluor-488 labeled) was polymerized in the presence of 1 μM fascin in TIRF buffer. After 120 seconds, the reaction mixture was replaced with a solution containing 500 nM fascin, 1.3 μM Alexa-Fluor-488–actin and 300 nM cofilin. Although most parts of the fascin-crosslinked filaments were disassembled, short, bundled barbed-end fragments, mainly composed of ATP- and ADP+Pi–actin persisted and continued to grow (arrowheads). Note that the fluorescence intensity of these growing bundles increased over time. Time is indicated in seconds. Scale bar: 5 μm. (D) Histogram of the correlation between cofilin decoration and the length of severed barbed end fragments of single filaments and fascin-crosslinked bundles. The inset shows representative bundled barbed-end fragments after cofilin addition. Values were obtained from experiments equivalent to those shown in A–C. *n=41, #n=44; &n=44; §n=153. (E) Left: as in C, except that 500 nM cofilin was added to the solution. The increase in fluorescence of the actin bundle after cofilin addition was quantified by densitometric analysis of the region in the red box. Right: the plot of fluorescence versus time of the boxed region demonstrates fluorescence increase after cofilin addition (arrow). (F) Cofilin increases the fluorescence of fascin-formed actin bundles in a concentration-dependent manner. The number of filaments per bundle after 10 minutes was estimated by assuming a linear relationship of fluorescence intensity and filament number as shown in Fig. 1B. For each condition, at least 30 bundles were analyzed. Boxes indicate 25th percentile, median and 75th percentile of all values; error bars indicate 10th and 90th percentile; minimal and maximal fluorescence intensities are also indicated. (G) Analyses of the binding of 1 μM cofilin to 1 μM F-actin (15% pyrene labeled) crosslinked by different amounts of fascin in KMEI buffer. Cofilin binding was slowed down by increasing amounts of fascin as shown by the enhanced half-time of the binding reaction. The line is a manual fit of the data points.

Fig. 2.

Cofilin-mediated severing of individual filaments and fascincrosslinked bundles. (A) 1.3 μM actin (30% Alexa-Fluor-488 labeled) was polymerized in the presence of 300 nM cofilin in TIRF buffer. The time-lapse micrographs show frequent severing of actin filaments (arrowheads). Time is indicated in seconds. (B) Left: direct visualization of EGFP–cofilin (green) binding to polymerizing Alexa-Fluor-633-labeled actin filaments (30% labeled; red). EGFP–cofilin preferentially bound to and decorated aged actin filaments, resulting in severing (arrowheads), whereas the barbed-end region was neither markedly decorated with EGFP–cofilin nor severed (asterisks). Time is indicated in seconds. Scale bar: 10 μm (A,B). Right: quantification of severing by cofilin and EGFP–cofilin. (C) Cofilin-mediated severing of fascin-bundled filaments. 1.3 μM actin (30% Alexa-Fluor-488 labeled) was polymerized in the presence of 1 μM fascin in TIRF buffer. After 120 seconds, the reaction mixture was replaced with a solution containing 500 nM fascin, 1.3 μM Alexa-Fluor-488–actin and 300 nM cofilin. Although most parts of the fascin-crosslinked filaments were disassembled, short, bundled barbed-end fragments, mainly composed of ATP- and ADP+Pi–actin persisted and continued to grow (arrowheads). Note that the fluorescence intensity of these growing bundles increased over time. Time is indicated in seconds. Scale bar: 5 μm. (D) Histogram of the correlation between cofilin decoration and the length of severed barbed end fragments of single filaments and fascin-crosslinked bundles. The inset shows representative bundled barbed-end fragments after cofilin addition. Values were obtained from experiments equivalent to those shown in A–C. *n=41, #n=44; &n=44; §n=153. (E) Left: as in C, except that 500 nM cofilin was added to the solution. The increase in fluorescence of the actin bundle after cofilin addition was quantified by densitometric analysis of the region in the red box. Right: the plot of fluorescence versus time of the boxed region demonstrates fluorescence increase after cofilin addition (arrow). (F) Cofilin increases the fluorescence of fascin-formed actin bundles in a concentration-dependent manner. The number of filaments per bundle after 10 minutes was estimated by assuming a linear relationship of fluorescence intensity and filament number as shown in Fig. 1B. For each condition, at least 30 bundles were analyzed. Boxes indicate 25th percentile, median and 75th percentile of all values; error bars indicate 10th and 90th percentile; minimal and maximal fluorescence intensities are also indicated. (G) Analyses of the binding of 1 μM cofilin to 1 μM F-actin (15% pyrene labeled) crosslinked by different amounts of fascin in KMEI buffer. Cofilin binding was slowed down by increasing amounts of fascin as shown by the enhanced half-time of the binding reaction. The line is a manual fit of the data points.

Fascin enhances cofilin-mediated severing both during polymerization and depolymerization

We reasoned that the increased actin fluorescence within growing bundles in the TIRF polymerization assays resulted from the formation of new barbed ends generated by cofilin severing, and that severing might be enhanced in fascin-crosslinked bundles when compared with single filaments. To test for barbed-end formation in an alternative, more quantitative fashion, pyrene assays were employed. Cofilin was previously shown to enhance actin polymerization in these assays (Carlier et al., 1997), possibly by filament severing, thereby creating new barbed ends for subsequent elongation (Carlier et al., 1997; Du and Frieden, 1998; Gandhi et al., 2009). Cofilin increased the rate of actin polymerization in a concentration-dependent manner, and 600 nM cofilin enhanced the rate of the assembly of 4 μM G-actin into F-actin approximately threefold (Fig. 3A,C). Fascin alone had virtually no effect on spontaneous actin assembly (Fig. 3B). However, the presence of fascin markedly enhanced the rate of cofilin-mediated actin polymerization, by as much as sevenfold, supporting our previous hypothesis of enhanced severing and barbed-end formation by cofilin in fascin-crosslinked bundles (Fig. 3B,C).

To evaluate filament depolymerization by cofilin, we monitored the spontaneous disassembly of 100 nM F-actin in KMEI buffer by measuring the decrease in pyrene fluorescence upon dilution. We found that the depolymerization rate was enhanced by cofilin alone in a concentration-dependent manner, by as much as fivefold (Fig. 3D–F). These effects of cofilin on spontaneous filament disassembly are in good agreement with results obtained in previous studies (Moseley et al., 2006; Gandhi et al., 2009). In contrast to non-bundled actin filaments, fascincrosslinked filaments depolymerized much slower (Fig. 3D–F). The fascin-dependent decrease in depolymerization rates is most probably caused by crosslinking of actin subunits and thus impaired monomer release from filament ends (Fig. 3D,E,G) (Schmoller et al., 2011). However, cofilin enhanced the depolymerization rate of fascin-bundled filaments in a concentration-dependent manner, by as much as 35-fold when compared with the respective controls without cofilin (Fig. 3F). This suggests that cofilin partially counteracted the inhibitory effect of fascin on actin disassembly by creating new filament ends in fascin-crosslinked bundles more efficiently than in isolated filaments (Fig. 3G). Because the rates of depolymerization, in contrast to polymerization, depends on both the number of barbed ends and the amount of crosslinking protein within the bundle, the enhancement of depolymerization rates by cofilin did not simply increase with higher fascin concentrations. Instead, it was maximal at a concentration of 20 nM fascin in the reaction, but decreased again at higher fascin concentrations (Fig. 3F). Notwithstanding this, the enhancement of depolymerization of fascin-bundled actin filaments by cofilin was higher when compared with non-bundled filaments for the entire range of fascin concentrations tested (Fig. 3F). Remarkably therefore, in spite of reduced cofilin binding kinetics in the presence of fascin (Fig. 2E), the latter was capable of enhancing cofilin activity to effect actin filament severing and disassembly in this assay.

Cofilin accumulates in retracting filopodia

Previous in vivo studies in B16-F1 and MTLn3 cells have shown that cofilin readily associates with the entire lamellipodium (Lai et al., 2008), and it probably modulates lamellipodium architecture and treadmilling by severing and depolymerizing aged actin filaments (Svitkina and Borisy, 1999; Iwasa and Mullins, 2007; Bugyi and Carlier, 2010). Another model suggests a cooperation of cofilin-induced barbed-end generation and Arp2/3-complex-mediated nucleation to drive lamellipodial actin assembly (Oser and Condeelis, 2009). Irrespective of the precise molecular mechanism, localization and functional interference studies commonly suggest a positive regulatory function for cofilin in driving lamellipodium protrusion and membrane ruffling (Mouneimne et al., 2004; Hotulainen et al., 2005; Iwasa and Mullins, 2007; Lai et al., 2008).

Inspired by our in vitro observations, we decided to reinvestigate the localization and dynamics of cofilin in migrating melanoma cells and fibroblasts. A polyclonal antibody was employed to determine the subcellular localization of endogenous cofilin in B16-F1 cells by indirect immunofluorescence. Consistent with previous findings, cofilin was found to accumulate in the lamellipodial actin network of migrating cells, indicating specificity of the antibody (Fig. 4A). Notably, however, cofilin was also found to be markedly enriched at the tips of some filopodia, while it was clearly absent from others (Fig. 4A). The localization of endogenous cofilin to filopodial tips was also confirmed in fixed T101/2 fibroblasts (Fig. 4B), suggesting that cofilin could affect filopodia dynamics in different cell types. In order to explore in more detail the puzzling finding that cofilin accumulates only in a subpopulation of filopodia, we reinvestigated the dynamics of fluorescently tagged cofilin in B16-F1 mouse melanoma cells. Interestingly, we found that mCherry–cofilin was weakly associated with the base of protruding filopodia (Fig. 5A), which is in line with proposed models in which filopodium formation is regulated by actin assembly by proteins of the filopodium tip complex (FTP) and disassembly by cofilin in the rear (Mattila and Lappalainen, 2008; Faix et al., 2009; Hotulainen and Hoogenraad, 2010). However, mCherry–cofilin changed its localization pattern significantly upon cessation of filopodium protrusion, now more prominently localizing along the entire shaft and/or massively accumulating in the filopodium tip (Fig. 5A; supplementary material Movie 9). Most notably, the localization of cofilin to the filopodium tip was exclusively detected in retracting filopodia, whereas the tips of protruding filopodia were virtually devoid of cofilin (Fig. 5B). Identical results were obtained with EGFP-tagged cofilin used previously (data not shown) (Lai et al., 2008). Since our in vitro results indicated a functional cooperation between cofilin and fascin, a well-established filopodial regulator, we also compared the dynamics of both proteins in vivo. To do this, we coexpressed EGFP-tagged fascin and mCherry–cofilin to analyze their relative dynamics in protruding and retracting filopodia. As expected, fascin was highly enriched in the entire shaft of protruding filopodia, whereas cofilin was, at best, weakly associated with the rear of these filopodia (Fig. 6; supplementary material Movie 10). However, upon cessation of protrusion, cofilin and fascin briefly colocalized in the shaft, followed by a decrease in fascin and overall increase in cofilin intensities, frequently culminating in strong accumulation of the latter in retracting filopodia tips (Fig. 6; supplementary material Movie 11). These data are consistent with the absence of cofilin from freshly polymerized barbed ends of filaments bundled by fascin in vitro (supplementary material Movie 8), and identify cofilin as a potential decisive factor in actin filament disassembly causing retraction of filopodial bundles in vivo. Consistent with this view, comparison of actin (mCherry) and cofilin (EGFP) dynamics in the same cells revealed that cofilin accumulation in filopodia upon cessation of protrusion and/or retraction directly correlates with a reduction of actin intensities, as expected if cofilin was to operate in actin filament disassembly at these sites (Fig. 7; supplementary material Movie 12). To study this directly, we used correlated live cell imaging and electron tomography to analyze the structure of filopodia captured by fixation in various phases of protrusion or withdrawal. Frequently, filopodia and microspikes in B16-F1 melanoma cells tagged with EGFP–fascin, performed translational and folding movements, including fragmentation and entry into the lamella, as previously described (Nemethova et al., 2008: supplementary material Movie 13). Additionally, however, filopodia labeled with cofilin showed characteristic retraction and kinking motions (Fig. 8A). Electron tomography revealed a correlation of the cofilin label with a marked fragmentation of actin filaments within the filopodium shaft, including the tip region (Fig. 8C,D). By contrast, neighboring filopodia in phases of protrusion that typically lacked the cofilin label showed intact filaments, arranged in parallel from base to tip (Fig. 8B).

Fig. 3.

Fascin enhances cofilin severing in a synergistic fashion. (A) Cofilin enhances actin assembly by creating new barbed ends. 4 μM G-actin (10% pyrene labeled) was polymerized in KMEI buffer in the presence of different amounts of cofilin. (B) Fascin enhances cofilin-mediated actin polymerization. 4 μM G-actin (10% pyrene labeled) was polymerized in KMEI buffer in the presence of 600 nM cofilin and increasing amounts of fascin, resulting in a dose-dependent acceleration of actin polymerization. (C) Comparison of polymerization rates mediated by different cofilin concentrations in the presence and absence of fascin. The relative polymerization rates correspond to the maximal slopes of the polymerization reactions shown in A and B. (D) Effects of cofilin and fascin on dilution-induced depolymerization of F-actin. 1 μM F-actin (30% pyrene labeled) either untreated or crosslinked by 0.2 and 1 μM fascin, respectively, was diluted to 100 nM in KMEI buffer with the cofilin concentrations indicated, and spontaneous actin disassembly followed, as measured by pyrene fluorescence. (E) Depolymerization rates of F-actin crosslinked with fascin at the concentrations indicated at different cofilin concentrations. Rates were obtained by measuring the initial slopes of depolymerization reactions as shown in D. (F) Comparison of the enhancement of depolymerization of actin filaments and fascin bundles by cofilin. The fold increase in actin disassembly was obtained by normalizing the depolymerization rates to the respective control experiments without cofilin. (G) Scheme depicting the differential influence of fascin and cofilin on filament depolymerization. (Top) Dilution-induced release of monomers from actin filaments (black arrows) occurs mainly at the barbed end. (Middle) Fascin probably inhibits filament disassembly by crosslinking terminal subunits, thus impairing dissociation of monomers (red arrow). (Bottom) Cofilin antagonizes fascin-mediated resistance to disassembly by creating new filament ends.

Fig. 3.

Fascin enhances cofilin severing in a synergistic fashion. (A) Cofilin enhances actin assembly by creating new barbed ends. 4 μM G-actin (10% pyrene labeled) was polymerized in KMEI buffer in the presence of different amounts of cofilin. (B) Fascin enhances cofilin-mediated actin polymerization. 4 μM G-actin (10% pyrene labeled) was polymerized in KMEI buffer in the presence of 600 nM cofilin and increasing amounts of fascin, resulting in a dose-dependent acceleration of actin polymerization. (C) Comparison of polymerization rates mediated by different cofilin concentrations in the presence and absence of fascin. The relative polymerization rates correspond to the maximal slopes of the polymerization reactions shown in A and B. (D) Effects of cofilin and fascin on dilution-induced depolymerization of F-actin. 1 μM F-actin (30% pyrene labeled) either untreated or crosslinked by 0.2 and 1 μM fascin, respectively, was diluted to 100 nM in KMEI buffer with the cofilin concentrations indicated, and spontaneous actin disassembly followed, as measured by pyrene fluorescence. (E) Depolymerization rates of F-actin crosslinked with fascin at the concentrations indicated at different cofilin concentrations. Rates were obtained by measuring the initial slopes of depolymerization reactions as shown in D. (F) Comparison of the enhancement of depolymerization of actin filaments and fascin bundles by cofilin. The fold increase in actin disassembly was obtained by normalizing the depolymerization rates to the respective control experiments without cofilin. (G) Scheme depicting the differential influence of fascin and cofilin on filament depolymerization. (Top) Dilution-induced release of monomers from actin filaments (black arrows) occurs mainly at the barbed end. (Middle) Fascin probably inhibits filament disassembly by crosslinking terminal subunits, thus impairing dissociation of monomers (red arrow). (Bottom) Cofilin antagonizes fascin-mediated resistance to disassembly by creating new filament ends.

Fig. 4.

Endogenous cofilin can accumulate in filopodia in different cell types. (A) B16-F1 melanoma cells cultivated on laminin-coated glass coverslips were fixed and labeled with cofilin antibody (green) and filamentous actin with Rhodamine-phalloidin (red). (Top row) Low magnification shows that cofilin accumulates in the entire lamellipodium. (Lower rows) Examples of cofilin enrichment in a subset of filopodia at higher magnification. (B) Subcellular localization of cofilin in T101/2 fibroblasts. (Top row) Cofilin accumulates in actin-rich membrane ruffles at many, individual cellular protrusions. (Bottom row) Cofilin also localizes to the distal tips of filopodial actin filaments in this cell type. The cells were labeled as described for A. All images are three-dimensional reconstructions from confocal sections. Scale bars: 10 μm.

Fig. 4.

Endogenous cofilin can accumulate in filopodia in different cell types. (A) B16-F1 melanoma cells cultivated on laminin-coated glass coverslips were fixed and labeled with cofilin antibody (green) and filamentous actin with Rhodamine-phalloidin (red). (Top row) Low magnification shows that cofilin accumulates in the entire lamellipodium. (Lower rows) Examples of cofilin enrichment in a subset of filopodia at higher magnification. (B) Subcellular localization of cofilin in T101/2 fibroblasts. (Top row) Cofilin accumulates in actin-rich membrane ruffles at many, individual cellular protrusions. (Bottom row) Cofilin also localizes to the distal tips of filopodial actin filaments in this cell type. The cells were labeled as described for A. All images are three-dimensional reconstructions from confocal sections. Scale bars: 10 μm.

Fig. 5.

Cofilin dynamics in protruding and retracting filopodia. (A) mCherry–cofilin expression pattern in a motile B16-F1 cell. During protrusion (upper panels), cofilin only weakly localizes to the base of filopodia and is absent in their tips. Upon retraction (lower panels), cofilin markedly accumulates in the tip region of these structures (arrowheads). Scale bar: 5 μm. (B) Quantification of protruding and retracting filopodia with cofilin accumulated in the tip. Note that cofilin accumulation is exclusively found in retracting filopodia.

Fig. 5.

Cofilin dynamics in protruding and retracting filopodia. (A) mCherry–cofilin expression pattern in a motile B16-F1 cell. During protrusion (upper panels), cofilin only weakly localizes to the base of filopodia and is absent in their tips. Upon retraction (lower panels), cofilin markedly accumulates in the tip region of these structures (arrowheads). Scale bar: 5 μm. (B) Quantification of protruding and retracting filopodia with cofilin accumulated in the tip. Note that cofilin accumulation is exclusively found in retracting filopodia.

Fig. 6.

Fascin and cofilin dynamics during protrusion and retraction of filopodia. EGFP–fascin and mCherry–cofilin were coexpressed in B16-F1 cells. Although cofilin is nearly absent in the protruding filopodium, it localizes to the shaft upon cessation of protrusion and accumulates in the tip region upon retraction. Fascin localization in the shaft is prominent during protrusion and decreases upon retraction. Note that fascin can still be found in the tip region of a retracting filopodium. Scale bar: 2 μm. Images were taken at 15-second intervals, and time is given in seconds.

Fig. 6.

Fascin and cofilin dynamics during protrusion and retraction of filopodia. EGFP–fascin and mCherry–cofilin were coexpressed in B16-F1 cells. Although cofilin is nearly absent in the protruding filopodium, it localizes to the shaft upon cessation of protrusion and accumulates in the tip region upon retraction. Fascin localization in the shaft is prominent during protrusion and decreases upon retraction. Note that fascin can still be found in the tip region of a retracting filopodium. Scale bar: 2 μm. Images were taken at 15-second intervals, and time is given in seconds.

Discussion

Cofilin was previously shown to cooperatively bind ADP–actin filaments, thereby enhancing their flexibility and the twist of the filament that eventually leads to filament breaking (McGough et al., 1997; Ressad et al., 1998; Blanchoin and Pollard, 1999; Pavlov et al., 2007; McCullough et al., 2008; Suarez ez al., 2011). However, the effects of actin crosslinking proteins on cofilin activity have not been investigated.

Cofilin-mediated severing of fascin-crosslinked actin bundles

By employing in vitro TIRF microscopy, we were able to show, for the first time, the dynamics of fascin-meditated fusion of single filaments into bundles, and directly demonstrate that fascin-mediated actin filament bundling is rapid, zippering together actin filaments at rates of up to 1000 subunits/second. We also confirmed that bundles formed by fascin are parallel, as reported previously (Ishikawa et al., 2003). Because the rate of spontaneous actin assembly into filaments is approximately two orders of magnitude slower than fascin-mediated bundling, we propose that filaments that grow in fascin–actin bundles are crosslinked immediately in vitro and probably also in vivo. We also used this in vitro assay to analyze the effects of cofilin on polymerizing, fascin-crosslinked actin bundles. Surprisingly, although we expected to observe reduced activities of cofilin on bundled filaments, as previously reported for spontaneously formed bundles (Michelot et al., 2007), we found that filament severing was greatly enhanced within these bundles. This was true in spite of slower binding kinetics of cofilin to fascin-bundled actin filaments as compared with isolated actin filaments, resulting in a dramatic increase in the number of barbed ends upon initial binding of cofilin. The enhancement of filament severing was proportional to the amount of both fascin and cofilin in the reaction mixture. Most notably, cofilin already efficiently severed filaments in bundles at concentrations that were not sufficient to sever individual actin filaments. How can these results be explained? It was proposed that the severing mechanism of cofilin relies on local changes of the helical twist of the filament upon cofilin binding. The F-actin twist was shown to be variable within a given filament: while the largest part of the filament twists by 167°, some segments twist only by 162° (Galkin et al., 2001; Galkin et al., 2003). Binding of cofilin to the latter shifts this equilibrium and induces 162° twists also in adjacent segments, which in turn promotes cooperative binding of additional cofilin molecules. However, efficient severing occurs predominantly at the borders of decorated and non-decorated segments of the actin filament (McGough et al., 1997; Prochniewicz et al., 2005; McCullough et al., 2008; De La Cruz and Sept, 2010, Suarez et al., 2011), so rapid decoration of the filament with cofilin will reduce severing frequency.

Fig. 7.

Comparison of cofilin and actin dynamics during filopodia protrusion and retraction. EGFP–cofilin and mCherry–actin were coexpressed in B16-F1 cells and imaged as indicated. Note the strong actin staining during filopodium protrusion (early time points), and its continuous decrease during cofilin accumulation and cessation of filopodium protrusion and retraction. Scale bar: 3 μm. Images were taken at 15-second intervals, and time is given in seconds.

Fig. 7.

Comparison of cofilin and actin dynamics during filopodia protrusion and retraction. EGFP–cofilin and mCherry–actin were coexpressed in B16-F1 cells and imaged as indicated. Note the strong actin staining during filopodium protrusion (early time points), and its continuous decrease during cofilin accumulation and cessation of filopodium protrusion and retraction. Scale bar: 3 μm. Images were taken at 15-second intervals, and time is given in seconds.

Based on these previous findings, two, not necessarily exclusive, mechanisms could account for the observed enhancement of cofilin severing by fascin. Crosslinking probably reduces the flexibility of the actin filament within the bundle. As a consequence, an actin filament segment cannot compensate for cofilin-induced twisting by relaxation of the entire filament (Fig. 9A). Thus, the mechanical stress generated upon cofilin binding would be restricted to the region between the crosslinkers, leading to rapid filament breaking (Fig. 9B). Alternatively, or in addition, fascin-mediated crosslinking could also counteract cooperative binding by cofilin (Fig. 9B). In this scenario, fascin crosslinking would prevent twisting of adjacent segments of the filament upon cofilin binding, leading to non-uniform decoration of the filament with cofilin, accompanied by more efficient severing (De La Cruz and Sept, 2010). The latter scenario is in line with our observation that cofilin binds to fascin-crosslinked bundles with slower kinetics than to isolated filaments. Moreover, fascin was recently shown to induce overtwisting of actin filaments upon bundling (Claessens et al., 2008). This effect might additionally account for an enhanced severing activity of cofilin due to larger differences in the twists of cofilin-bound compared with fascin-bound actin filament segments. Irrespective of this, the efficiency of cofilin severing was enhanced by increasing concentrations of fascin in the reaction, despite reduced cofilin binding kinetics, and filament severing was already observable at concentrations of cofilin too low to induce marked severing of isolated filaments, which is consistent with both models.

Fig. 8.

Correlated live cell imaging and electron tomography of protruding and retracting filopodia. (A) Time-lapse series (every second frame) of B16-F1 melanoma cell transfected with EGFP–cofilin and mCherry–fascin. Time between the frames shown was 30 seconds. b, c, d indicate individual filopodia that were protruding (b) or retracting (c,d) at the time of fixation after last frame of fluorescence imaging. The final panel shows the fixed cell under phase-contrast optics. Cofilin is present along the retracting filopodium shaft (d) or at the tip (c). (B,C,D) Tomogram sections of the corresponding filopodia in A. The regions indicated by brackets are enlarged in B′,C′ and D′. For technical reasons, the zero tilt image of filopodium ‘c’ is shown in C and C′ (lower panel). Scale bars: 5 μm (A); 250 nm (B–D); 100 nm (B′–D′).

Fig. 8.

Correlated live cell imaging and electron tomography of protruding and retracting filopodia. (A) Time-lapse series (every second frame) of B16-F1 melanoma cell transfected with EGFP–cofilin and mCherry–fascin. Time between the frames shown was 30 seconds. b, c, d indicate individual filopodia that were protruding (b) or retracting (c,d) at the time of fixation after last frame of fluorescence imaging. The final panel shows the fixed cell under phase-contrast optics. Cofilin is present along the retracting filopodium shaft (d) or at the tip (c). (B,C,D) Tomogram sections of the corresponding filopodia in A. The regions indicated by brackets are enlarged in B′,C′ and D′. For technical reasons, the zero tilt image of filopodium ‘c’ is shown in C and C′ (lower panel). Scale bars: 5 μm (A); 250 nm (B–D); 100 nm (B′–D′).

The physiological relevance of the concerted activities of fascin and cofilin

It is well established that cofilin regulates the protrusion of lamellipodia, in which it is strongly enriched (Lai et al., 2008; Oser and Condeelis, 2009). Also, its knockdown was observed to reduce the actin filament turnover in lamellipodia and other actin structures such as stress fibers (Hotulainen et al., 2005). This effect was interpreted as F-actin stabilization due to reduced depolymerization, and hence depletion of the actin monomer pool (Kiuchi et al., 2007).

We show here, for the first time, that cofilin is also highly enriched in the tip region of retracting filopodia and that specific localization patterns of fascin and cofilin are directly correlated with filopodial protrusion, stalling and retraction. Interestingly, the transient association of cofilin with retracting rather than protruding filopodia suggests it specifically operates in promoting the severing and disassembly of actin filaments in these structures, as in lamellipodia (Hotulainen et al., 2005; Lai et al., 2008). This notion is supported by our analyses of actin filament organization in filopodia at different stages of protrusion and retraction using a combination of live cell imaging and electron tomography. Our experiments revealed compact bundles of actin filaments in filopodia during protrusion, as expected. However, filopodia at different stages of retraction had regions of fragmented actin filaments that precisely correlated with the accumulation of cofilin, established by preceding live-cell imaging. This criss-cross arrangement of short filaments is reminiscent of the actin organization reported for Dictyostelium filopodia, presumably fixed during their retraction (Medalia et al., 2007), indicating that this mechanism of filopodium retraction might have been conserved throughout evolution. Furthermore, enhancement of cofilin-induced severing by fascin, as observed in vitro, would be consistent with a transient colocalization of both proteins in filopodia (Figs 6 and 8C). However, the frequently observed strong accumulation of cofilin in the tip region of the filopodium–a region assumed to largely contain filament barbed ends–is striking. The occurrence of rearward moving cofilin puncta within filopodia (supplementary material Movies 9 and 10) indicates continuous actin polymerization in the filopodium tip even when net elongation stalls, suggesting that cofilin might bind to newly polymerized actin filaments. Taking into account that cofilin preferentially binds to ADP–F-actin, it is possible that phosphate release is enhanced by components of the filopodium tip complex. Importantly, proteins of the formin and Ena/VASP family, which are crucial components of the filopodium tip complex (Schirenbeck et al., 2005, Kwiatkowski et al., 2007; Breitsprecher et al., 2008,) were suggested to mediate actin filament elongation by coupling ATP hydrolysis and phosphate release (Dickinson and Purich, 2002; Dickinson et al., 2004; Romero et al., 2004; Romero et al., 2007). Thus, these proteins might not only be important players in the initiation and elongation of filopodia but could also operate in the regulation of filopodium length and lifetime by providing ADP–F-actin as a depolymerizable substrate for cofilin; a hypothesis deserving future investigation. However, it should be emphasized that proper, subcellular cofilin positioning must rely on additional components, because in these cells cofilin is actively kept away from actin filaments in structures other than lamellipodia and retracting filopodia, so the preference for ADP–actin filaments might be one of many aspects contributing to cofilin localization and dynamics in vivo.

Fig. 9.

Proposed mechanism for cofilin-mediated severing of fascincrosslinked filaments. (A) Single actin filaments change their torsional twist upon cofilin (blue circles) binding, resulting in complete decoration of the filament in a cooperative manner. Severing events occur stochastically. Red squares mark sites of severing. (B) Fascin (green bars)-crosslinked filaments are less flexible than free filaments and might therefore be impaired in relaxation upon cofilin binding, resulting in enhanced mechanical stress that leads to enhanced filament severing. Alternatively or in addition, fascin might also impair cooperative binding by cofilin, resulting in a discontinuous decoration of the filament in the bundle and enhanced cofilin severing activity. (C) A model of the potential role of cofilin in filopodium disassembly. During protrusion, filaments are elongated by proteins of the filopodium tip complex (FTP) and immediately crosslinked by fascin, while cofilin is largely absent. Upon cessation of protrusion, cofilin targets to the shaft and strongly accumulates in the tip region of the filopodium by an unknown mechanism, and promotes the disassembly of filopodial actin filaments. Eventually, cofilin-mediated filament severing leads to retraction of the filopodium into the cell body. Upwards and downwards arrow indicates protrusion and retraction, respectively.

Fig. 9.

Proposed mechanism for cofilin-mediated severing of fascincrosslinked filaments. (A) Single actin filaments change their torsional twist upon cofilin (blue circles) binding, resulting in complete decoration of the filament in a cooperative manner. Severing events occur stochastically. Red squares mark sites of severing. (B) Fascin (green bars)-crosslinked filaments are less flexible than free filaments and might therefore be impaired in relaxation upon cofilin binding, resulting in enhanced mechanical stress that leads to enhanced filament severing. Alternatively or in addition, fascin might also impair cooperative binding by cofilin, resulting in a discontinuous decoration of the filament in the bundle and enhanced cofilin severing activity. (C) A model of the potential role of cofilin in filopodium disassembly. During protrusion, filaments are elongated by proteins of the filopodium tip complex (FTP) and immediately crosslinked by fascin, while cofilin is largely absent. Upon cessation of protrusion, cofilin targets to the shaft and strongly accumulates in the tip region of the filopodium by an unknown mechanism, and promotes the disassembly of filopodial actin filaments. Eventually, cofilin-mediated filament severing leads to retraction of the filopodium into the cell body. Upwards and downwards arrow indicates protrusion and retraction, respectively.

In contrast to promoting actin filament disassembly, cofilin has also been proposed to drive the assembly of actin filaments, at high concentrations both in vitro (Andrianantoandro and Pollard, 2006) and in vivo (Oser et al., 2009), although our own results comparing actin and cofilin dynamics in the lamellipodia of different cell types were not confirmatory of this view (Lai et al., 2008). However, such a function of cofilin cannot be excluded for other subcellular locations, e.g. invadopodia and matrix-degrading adhesive structures employed by cancer cells during metastasis. Interestingly, both fascin and cofilin are key components of invadopodia and are crucial for matrix invasion. Both proteins localize to the actin-rich region of these structures, and the depletion of either fascin or cofilin results in decreased invadopodia lifetimes and lower actin levels (Yamaguchi et al., 2005; Oser et al., 2009; Li et al., 2010). Considering the cooperation between cofilin and fascin in filament severing observed in vitro, it is tempting to speculate that their combined activities might result in distinct outcomes in different conditions or subcellular structures, e.g. driving actin polymerization in invadopodia while disassembling actin filaments in filopodia. Our data add to the list of functions established for cofilin in live cells, by demonstrating its transient engagement in filopodia retraction.

Materials and Methods

Constructs and protein purification

Murine non-muscle cofilin-1 (also known as n-cofilin) and human EGFP-tagged n-cofilin were purified as described previously (Lai et al., 2008). mCherry–actin (Koestler et al., 2008), EGFP–fascin (Adams and Schwartz, 2000) and EGFP–cofilin (Mannherz et al., 2005) were described previously. mCherry–cofilin was produced by excision from pEGFP–cofilin with NheI and BsrGI and ligation into the mCherry vector at the corresponding sites. Human fascin was amplified from a human macrophage cDNA library (Invitrogen) and inserted into the BamHI and XhoI sites of pGEX-6P1. Briefly, GST-tagged cofilins and GST-tagged fascin were expressed in Escherichia coli strain Rosetta (Promega) and purified from bacterial extracts on glutathione-conjugated agarose (Sigma-Aldrich, Germany) using standard procedures. The GST tags were cleaved by incubating the purified fusion proteins with PreScission protease (GE Heathcare) in phosphate-buffered saline (PBS), pH 7.3, supplemented with 1 mM dithiothreitol (DTT) and 1 mM EDTA overnight at 4°C. After cleavage, the GST tags were removed by gel filtration on a 26/60 Superdex G75 column (GE Healthcare). Cofilin- or fascin-containing fractions were pooled, dialyzed against 200 mM KCl, 1 mM DTT, 60% glycerol and 20 mM imidazole (pH 7.4) and stored at −23°C. Ca2+-ATP–actin was purified from rabbit skeletal muscle according to the method of Spudich and Watt (Spudich and Watt, 1971), stored in G-buffer (5 mM Tris-HCl pH 8.0 and 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ATP) and prepared for time-lapse total internal reflection fluorescence microscopy (TIRFM) and pyrene assays as described previously (Kuhn and Pollard, 2005; Schirenbeck et al., 2006). For two-color TIRF experiments, rabbit muscle actin was labeled on Cys374 with Alexa-Fluor-633–maleimide (Invitrogen). Protein concentrations were determined by absorption spectroscopy using extinction coefficients predicted from the amino acid sequences using Vector NTI software (Invitrogen).

In vitro TIRF microscopy

TIRFM with Alexa-Fluor-488-labeled actin was performed essentially as described previously (Breitsprecher et al., 2011). Images from an Olympus IX-81 inverted microscope were captured every 5 or 10 seconds using a Hamamatsu Orca-R2 CCD camera (Hamamatsu Corp., Bridgewater, NJ). For two-color TIRFM experiments, Alexa-Fluor-633-labeled actin was excited at 632 nm with a 30 mW HeNe laser (DS Uniphase Corp., Milpitas, CA) with exposure times of 300 mseconds, and EGFP–cofilin was excited at 488 nm with a 30 mW DPSS laser (Novalux, Sunnyvale, CA) with exposure times of 150 mseconds. The specimens were excited successively with time intervals <20 mseconds between the excitations to minimize time-shifts. The pixel size corresponded to 0.11 μm.

Reactions in TIRF assays contained 1.3 μM actin (labeled with 30% Alexa-Fluor-488 or -633). Before the experiments, Ca2+-ATP–actin was converted to Mg2+-ATP–actin by addition of 10× magnesium-exchange buffer (1 mM MgCl2, 10 mM EGTA, pH 7.4). Polymerization experiments were performed in TIRF buffer [10 mM imidazole, 50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 0.2 mM ATP, 10 mM DTT, 15 mM glucose, 20 μg/ml catalase, 100 μg/ml glucose oxidase, and 0.5% methylcellulose (4000 cP), pH 7.4]. The fluorescence intensity of bundles was analyzed using the line-scan tool in ImageJ software (http://rsbweb.nih.gov/ij/), and subsequently normalized to the fluorescence intensity of a single actin filament from the same experiment to calculate the number of filaments per bundle. Small filaments transiently interacting with the surface before their rapid disappearance were not included in the analyses.

Pyrene-actin assays

Before experiments, Ca2+-ATP–actin was converted to Mg2+-ATP–actin by addition of 10× Mg-exchange buffer. For assembly assays, dilutions of proteins to be assayed were prepared in protein storage buffer (200 mM KCl, 20 mM Hepes, 1 mM DTT, pH 7.3) and 10× low salt KMEI buffer (250 mM KCl, 10 mM MgCl2, 10 mM EGTA, 100 mM imidazole, pH 7.3) was added. Anti-foam 204 (Sigma) was added to the mixture to reach a final concentration of 0.005%. 180 μl aliquots were placed in an 8-well microtiter assembly strip (Thermo Scientific). 18 μl of a 40 μM solution of 10% pyrene-labeled G-actin (in 2 mM Tris-HCl, pH 8.0, 0.2 mM ATP, 0.1 mM CaCl2, 0.5 mM DTT) were placed in another 8-well microtiter assembly strip. The assembly reaction was initiated by transferring 162 μl of the protein solution to 18 μl of pyrene-labeled actin. The polymerization of actin was followed by measuring the fluorescence increase of pyrene-actin (excitation at 364 nm and emission at 407 nm) in a fluorescence plate reader for at least 1500 seconds.

Spontaneous depolymerization of F-actin was analyzed by diluting 10 μl of a 1 μM F-actin solution in 1× KMEI buffer (50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 10 mM imidazole, pH 7.3) crosslinked by 500 nM fascin into 90 μl 1× KMEI buffer containing the cofilin concentration as indicated. The decrease of pyrene fluorescence was monitored with a Jasco FP-6500 fluorimeter (excitation at 364 nm and emission at 407 nm), and the relative depolymerization rate was derived from the initial slope of the depolymerization reaction. Cofilin-binding to bundled F-actin was analyzed by adding 10 μl of a solution of 10 μM F-actin crosslinked with 0.15–10 μM fascin to 90 μl of a solution of 1.11 μM cofilin in 1× KMEI buffer. After rapid mixing, the decrease in pyrene fluorescence was monitored with a Jasco FP-6500 fluorimeter.

Low-speed sedimentation assays

G-actin (5 μM) were polymerized in 1× KMEI buffer in the presence of fascin and cofilin concentrations as indicated. Bundles were sedimented by centrifugation for 30 minutes at 11,000 g, and supernatants and pellets were analyzed by SDS-PAGE and Coomassie-Blue-staining. Densitometric analysis of band intensities was performed with ImageJ software (http://rsbweb.nih.gov/ij/).

Cells and transfections

Mouse embryonic Ch3T101/2 (T101/2) fibroblasts and B16-F1 mouse melanoma cells (ATTC CRL-6323) were grown in high glucose DMEM (Gibco) supplemented with 10% FCS (PAA Laboratories, Pasching, Austria) and 2 mM glutamine. Transfections were performed overnight using Superfect (Quiagen GmbH, Hilden, Germany) and FuGene 6 (Roche, Basel, Switzerland) according to manufacturers' instructions, respectively.

Microscopy

For indirect immunofluorescence microscopy, T101/2 fibroblasts were plated on uncoated glass coverslips, and B16-F1 cells were plated on acid-washed coverslips coated with 25 μg/ml laminin (Sigma). After fixation with 4% paraformaldehyde in PBS containing 2.7 mM KCl, 1.8 mM KH2PO4, 10 mM Na2HPO4, 140 mM NaCl, pH 7.3 for 15 minutes, the specimens were treated with PBS containing 100 mM glycine and 0.1% Triton X-100 for 15 minutes. After washing five times with PBS containing 0.05% (v/v) cold fish gelatin (Sigma) and 0.5% (w/v) bovine serum albumin (Sigma), the cells were incubated with a polyclonal anti-n-cofilin antibody (Dianova, Pinole, CA) at a dilution of 1:1000 in the same buffer overnight and then incubated with Alexa-Fluor-488-conjugated goat anti-rabbit IgG (Molecular Probes). F-actin was visualized with tetramethylrhodamine B isothiocyanate (TRITC)-conjugated phalloidin (Sigma). Confocal fluorescence images were recorded in multi-track mode with a Zeiss LSM 510 confocal microscope, equipped with a 63/1.3 Plan-Neofluar objective. Data were processed with Adobe Photoshop and CorelDraw software.

For live-cell imaging, B16-F1 cells were plated on acid-washed coverslips coated with 25 μg/ml laminin (Sigma), mounted in an open heating chamber (Warner Instruments, Reading, UK), and observed at 37°C with a Zeiss Axiovert S100TV inverted microscope (Carl Zeiss, Jena, Germany) equipped with a 63×/1.4 NA plan-apochromatic objective, and a cooled CCD camera (Coolsnap HQ2, Photometrics, Roper Scientific, Germany) with a filter wheel and shutters controlled by Metamorph software (Molecular Devices).

Correlated live-cell imaging and electron tomography were performed essentially as described previously (Auinger and Small, 2008; Urban et al., 2010). Briefly, B16 melanoma cells expressing mCherry–fascin and EGFP–cofilin were plated onto coverslips coated with a Formvar film embossed with grid patterns for cell relocation. Selected cells were imaged at 37°C in an open, temperature-controlled chamber using a 100×, 1.4 NA phase-contrast objective on an inverted Zeiss Observer fluorescence microscope equipped with an LED light source (CoolLed, Andover, UK) and a rear-illuminated cooled CCD Micromax camera (Roper Scientific). After recording paired images in two fluorescent channels every 15 seconds, the growth medium in the chamber was rapidly aspirated and replaced with the fixative mixture containing 0.5% Triton X-100 and 0.25% glutaraldehyde in cytoskeleton buffer (10 mM MES, 150 mM NaCl, 5 mM EGTA, 5 mM glucose, 5 mM MgCl2, pH 6.1). After 1 minute, the fixative mixture was exchanged for 2% glutaraldehyde in the same buffer supplemented with 10 μg/ml phalloidin and stored at 4°C in this solution until use. For electron microscopy, the Formvar film was peeled from the coverslip under cytoskeleton buffer, inverted onto the buffer surface and a 50 mesh EM grid positioned onto the region of the film containing the imaged cell using forceps mounted in a micromanipulator. The grids were stained with 7% sodium silicotungstate containing a gold sol, as described previously (Urban et al., 2010) and a tomogram tilt series recorded in an FEI Polara electron microscope at 300 kV using primary magnifications from 23–31,000× at a defocus of 5 μm. Tomograms were generated using IMOD software (Kremer et al., 1996).

Acknowledgements

We thank Annette Breskott and Brigitte Denker for excellent technical assistance. We also thank Josephine Adams (Cleveland, USA), Hans Georg Mannherz (Bochum, Germany) and Elena Korenbaum for kindly providing expression constructs and T101/2 cells.

Funding

This work was supported by the Deutsche Forschungsgemeinschaft [grant number FA 330/4-2 to J.F., grant number RO 2414/1-2 to K.R., grant number SPP1464 to J.V.S.]; the Austria Science Foundation [grant number FWF project P21292-B09]; and the Vienna Science Research and Technology Fund (WWTF).

References

Adams
J. C.
,
Schwartz
M. A.
(
2000
).
Stimulation of fascin spikes by thrombospondin-1 is mediated by the GTPases Rac and Cdc42
.
J. Cell Biol.
150
,
807
-
822
.
Auinger
S.
,
Small
J. V.
(
2008
).
Correlated light and electron microscopy of the cytoskeleton
.
Methods in Cell Biol.
88
,
257
-
272
.
Andrianantoandro
E.
,
Pollard
T. D.
(
2006
).
Mechanism of actin filament turnover by severing and nucleation at different concentrations of ADF/cofilin
.
Mol. Cell
24
,
13
-
23
.
Aratyn
Y. S.
,
Schaus
T. E.
,
Taylor
E. W.
,
Borisy
G. G.
(
2007
).
Intrinsic dynamic behavior of fascin in filopodia
.
Mol. Biol. Cell
18
,
3928
-
3940
.
Bernstein
B.W.
,
Bamburg
J. R.
(
2010
).
ADF/cofilin: a functional node in cell biology
.
Trends Cell Biol.
4
,
187
-
195
.
Blanchoin
L.
,
Pollard
T. D.
(
1999
).
Mechanism of interaction of Acanthamoeba actophorin (ADF/Cofilin) with actin filaments
.
J. Biol. Chem.
274
,
15538
-
15546
.
Bobkov
A. A.
,
Muhlrad
A.
,
Shvetsov
A.
,
Benchaar
S.
,
Scoville
D.
,
Almo
S. C.
,
Reisler
E.
(
2004
).
Cofilin (ADF) affects lateral contacts in F-actin
.
J. Mol. Biol.
337
,
93
-
104
.
Breitsprecher
D.
,
Kiesewetter
A. K.
,
Linkner
J.
,
Urbanke
C.
,
Resch
G. P.
,
Small
J. V.
,
Faix
J.
(
2008
).
Clustering of VASP actively drives processive, WH2 domain-mediated actin filament elongation
.
EMBO J.
27
,
2943
-
2954
.
Breitsprecher
D.
,
Kiesewetter
A. K.
,
Linkner
J.
,
Vinzenz
M.
,
Stradal
T. E. B.
,
Small
J. V.
,
Curth
U.
,
Dickinson
R. B.
,
Faix
J.
(
2011
).
Molecular mechanism of Ena/VASP-mediated actin-filament elongation
.
EMBO J.
30
,
456
-
467
.
Bugyi
B.
,
Carlier
M. F.
(
2010
).
Control of actin filament treadmilling in cell motility
.
Annu. Rev. Biophys.
39
,
449
-
470
.
Cao
W.
,
Goodarzi
J. P.
,
De La Cruz
E. M.
(
2006
).
Energetics and kinetics of cooperative cofilin-actin filament interactions
.
J. Mol. Biol.
361
,
257
-
267
.
Carlier
M. F.
,
Laurent
V.
,
Santolini
J.
,
Melki
R.
,
Didry
D.
,
Xia
G. X.
,
Hong
Y.
,
Chua
N. H.
,
Pantaloni
D.
(
1997
).
Actin depolymerizing factor (ADF/cofilin) enhances the rate of filament turnover: implication in actin-based motility
.
J. Cell Biol.
136
,
1307
-
1322
.
Carlier
M. F.
,
Pantaloni
D.
,
Evans
J. A.
,
Lambooy
P. K.
,
Korn
E. D.
,
Webb
M. R.
(
1988
).
The hydrolysis of ATP that accompanies actin polymerization is essentially irreversible
.
FEBS Lett.
235
,
211
-
214
.
Chesarone
M. A.
,
Goode
B. L.
(
2009
).
Actin nucleation and elongation factors: mechanisms and interplay
.
Curr. Opin. Cell Biol.
21
,
28
-
37
.
Claessens
M. M.
,
Semmrich
C.
,
Ramos
L.
,
Bausch
A. R.
(
2008
).
Helical twist controls the thickness of F-actin bundles
.
Proc. Natl. Acad. Sci.
USA105
,
8819
-
8822
.
Costantino
S.
,
Kent
C. B.
,
Godin
A. G.
,
Kennedy
T. E.
,
Wiseman
P. W.
,
Fournier
A. E.
(
2008
).
Semi-automated quantification of filopodial dynamics
.
J. Neurosci. Methods
171
,
165
-
173
.
Dent
E. W.
,
Kwiatkowski
A. V.
,
Mebane
L. M.
,
Philippar
U.
,
Barzik
M.
,
Rubinson
D. A.
,
Gupton
S.
,
Van Veen
J. E.
,
Furman
C.
,
Zhang
J.
,
Alberts
A. S.
,
Mori
S.
,
Gertler
F. B.
(
2007
).
Filopodia are required for cortical neurite initiation
.
Nat. Cell Biol.
9
,
1347
-
1359
.
De La Cruz
E. M.
(
2005
).
Cofilin binding to muscle and non-muscle actin filaments: isoform-dependent cooperative interactions
.
J. Mol. Biol.
346
,
557
-
564
.
De La Cruz
E. M.
,
Sept
D.
(
2010
).
The kinetics of cooperative cofilin binding reveals two states of the cofilin-actin filament
.
Biophys. J.
98
,
1893
-
1901
.
DeRosier
D. J.
,
Edds
K. T.
(
1980
).
Evidence for fascin cross-links between the actin filaments in coelomocyte filopodia
.
Exp. Cell Res.
126
,
490
-
494
.
Dickinson
R. B.
,
Purich
D. L.
(
2002
).
Clamped-filament elongation model for actin-based motors
.
Biophys. J.
82
,
605
-
617
.
Dickinson
R. B.
,
Caro
L.
,
Purich
D. L.
(
2004
).
Force generation by cytoskeletal filament end-tracking proteins
.
Biophys. J.
87
,
2838
-
2854
.
Du
J.
,
Frieden
C.
(
1998
).
Kinetic studies on the effect of yeast cofilin on yeast actin polymerization
.
Biochemistry
37
,
13276
-
13284
.
Faix
J.
,
Breitsprecher
D.
,
Stradal
T. E.
,
Rottner
K.
(
2009
).
Filopodia: Complex models for simple rods
.
Int. J. Biochem. Cell Biol.
41
,
1656
-
1664
.
Galkin
V. E.
,
Orlova
A.
,
Lukoyanova
N.
,
Wriggers
W.
,
Egelman
E. H.
(
2001
).
Actin depolymerizing factor stabilizes an existing state of F-actin and can change the tilt of F-actin subunits
.
J. Cell Biol.
153
,
75
-
86
.
Galkin
V. E.
,
Orlova
A.
,
VanLoock
M. S.
,
Shvetsov
A.
,
Reisler
E.
,
Egelman
E. H.
(
2003
).
ADF/cofilin use an intrinsic mode of F-actin instability to disrupt actin filaments
.
J. Cell Biol.
163
,
1057
-
1066
.
Gandhi
M.
,
Achard
V.
,
Blanchoin
L.
,
Goode
B. L.
(
2009
).
Coronin switches roles in actin disassembly depending on the nucleotide state of actin
.
Mol. Cell
34
,
364
-
374
.
Gehler
S.
,
Shaw
A. E.
,
Sarmiere
P. D.
,
Bamburg
J. R.
,
Letourneau
P. C.
(
2004
).
Brain-derived neurotrophic factor regulation of retinal growth cone filopodial dynamics is mediated through actin depolymerizing factor/cofilin
.
J. Neurosci.
24
,
10741
-
10749
.
Geraldo
S.
,
Gordon-Weeks
P. R.
(
2009
).
Cytoskeletal dynamics in growth-cone steering
.
J. Cell. Sci.
122
,
3595
-
3604
.
Hotulainen
P.
,
Hoogenraad
C. C.
(
2010
).
Actin in dendritic spines: connecting dynamics to function
.
J. Cell Biol.
189
,
619
-
629
.
Hotulainen
P.
,
Paunola
E.
,
Vartiainen
M. K.
,
Lappalainen
P.
(
2005
).
Actindepolymerizing factor and cofilin-1 play overlapping roles in promoting rapid F-actin depolymerization in mammalian nonmuscle cells
.
Mol. Biol. Cell
16
,
649
-
664
.
Insall
R. H.
,
Machesky
L. M.
(
2009
).
Actin dynamics at the leading edge: from simple machinery to complex networks
.
Dev. Cell.
17
,
310
-
322
.
Ishikawa
R.
,
Sakamoto
T.
,
Ando
T.
,
Higashi-Fujime
S.
,
Kohama
K.
(
2003
).
Polarized actin bundles formed by human fascin-1: their sliding and disassembly on myosin II and myosin V in vitro
.
J. Neurochem.
87
,
676
-
685
.
Iwasa
J. H.
,
Mullins
R. D.
(
2007
).
Spatial and temporal relationships between actin-filament nucleation, capping, and disassembly
.
Curr. Biol.
17
,
395
-
406
.
Kiuchi
T.
,
Ohashi
K.
,
Kurita
S.
,
Mizuno
K.
(
2007
).
Cofilin promotes stimulus-induced lamellipodium formation by generating an abundant supply of actin monomers
.
J. Cell Biol.
177
,
465
-
476
.
Koestler
S. A.
,
Auinger
S.
,
Vinzenz
M.
,
Rottner
K.
,
Small
J. V.
(
2008
).
Differentially oriented populations of actin filaments generated in lamellipodia collaborate in pushing and pausing at the cell front
.
Nat. Cell Biol.
10
,
306
-
313
.
Kuhn
J. R.
,
Pollard
T. D.
(
2005
).
Real-time measurements of actin filament polymerization by total internal reflection fluorescence microscopy
.
Biophys. J.
88
,
1387
-
1402
.
Kremer
J. R.
,
Mastronarde
D. N.
,
McIntosh
J. R.
(
1996
).
Computer visualization of threedimensional image data using IMOD
.
J. Struct. Biol.
116
,
71
76
.
Kwiatkowski
A. V.
,
Rubinson
D. A.
,
Dent
E. W.
,
Edward van Veen
J.
,
Leslie
J. D.
,
Zhang
J.
,
Mebane
L. M.
,
Philippar
U.
,
Pinheiro
E. M.
,
Burds
A. A.
,
Bronson
R. T.
,
Mori
S.
,
Fässler
R
,
Gertler
F. B.
(
2007
).
Ena/VASP Is Required for neuritogenesis in the developing cortex
.
Neuron
56
,
441
-
455
.
Lai
F. P.
,
Szczodrak
M.
,
Block
J.
,
Faix
J.
,
Breitsprecher
D.
,
Mannherz
H. G.
,
Stradal
T. E.
,
Dunn
G. A.
,
Small
J. V.
,
Rottner
K.
(
2008
).
Arp2/3 complex interactions and actin network turnover in lamellipodia
.
EMBO J.
27
,
982
-
992
.
Li
A.
,
Dawson
J. C.
,
Forero-Vargas
M.
,
Spence
H. J.
,
Yu
X.
,
Konig
I.
,
Anderson
K.
,
Machesky
L. M.
(
2010
).
The actin-bundling protein fascin stabilizes actin in invadopodia and potentiates protrusive invasion
.
Curr. Biol.
20
,
339
-
345
.
Lieleg
O.
,
Claessens
M. M.
,
Heussinger
C.
,
Frey
E.
,
Bausch
A. R.
(
2007
).
Mechanics of bundled semiflexible polymer networks
.
Phys. Rev. Lett.
99
,
088102
.
Loisel
T. P.
,
Boujemaa
R.
,
Pantaloni
D.
,
Carlier
M. F.
(
1999
).
Reconstitution of actin-based motility of Listeria and Shigella using pure proteins
.
Nature
401
,
613
-
616
.
Maciver
S. K.
,
Wachsstock
D. H.
,
Schwarz
W. H.
,
Pollard
T. D.
(
1991
).
The actin filament severing protein actophorin promotes the formation of rigid bundles of actin filaments crosslinked with α-actinin
.
J. Cell Biol.
115
,
1621
-
1628
.
Mannherz
H. G.
,
Gonsior
S. M.
,
Gremm
D.
,
Wu
X.
,
Pope
B. J.
,
Weeds
A. G.
(
2005
).
Activated cofilin colocalises with Arp2/3 complex in apoptotic blebs during programmed cell death
.
Eur. J. Cell Biol.
84
,
503
-
515
.
Mattila
P. K.
,
Lappalainen
P.
(
2008
).
Filopodia: molecular architecture and cellular functions
.
Nat. Rev. Mol. Cell Biol.
9
,
446
-
454
.
McCullough
B. R.
,
Blanchoin
L.
,
Martiel
J. L.
,
De la Cruz
E. M.
(
2008
).
Cofilin increases the bending flexibility of actin filaments: implications for severing and cell mechanics
.
J. Mol. Biol.
381
,
550
-
558
.
McGough
A.
,
Chiu
W.
(
1999
).
ADF/cofilin weakens lateral contacts in the actin filament
.
J. Mol. Biol.
291
,
513
-
519
.
McGough
A.
,
Pope
B.
,
Chiu
W.
,
Weeds
A.
(
1997
).
Cofilin changes the twist of F-actin: implications for actin filament dynamics and cellular function
.
J. Cell Biol.
138
,
771
-
781
.
Medalia
O.
,
Beck
M.
,
Ecke
M.
,
Weber
I.
,
Neujahr
R.
,
Baumeister
W.
,
Gerisch
G.
(
2007
).
Organization of actin networks in intact filopodia
.
Curr. Biol.
17
,
79
-
84
.
Michelot
A.
,
Berro
J.
,
Guerin
C.
,
Boujemaa-Paterski
R.
,
Staiger
C. J.
,
Martiel
J. L.
,
Blanchoin
L.
(
2007
).
Actin-filament stochastic dynamics mediated by ADF/cofilin
.
Curr. Biol.
17
,
825
-
833
.
Moseley
J. B.
,
Okada
K.
,
Balcer
H. I.
,
Kovar
D. R.
,
Pollard
T. D.
,
Goode
B. L.
(
2006
).
Twinfilin is an actin-filament-severing protein and promotes rapid turnover of actin structures in vivo
.
J. Cell. Sci.
119
,
1547
-
1557
.
Mouneimne
G.
,
Soon
L.
,
DesMarais
V.
,
Sidani
M.
,
Song
X.
,
Yip
S. C.
,
Ghosh
M.
,
Eddy
R.
,
Backer
J. M.
,
Condeelis
J.
(
2004
).
Phospholipase C and cofilin are required for carcinoma cell directionality in response to EGF stimulation
.
J. Cell Biol.
166
,
697
-
708
.
Nemethova
M.
,
Auinger
S.
,
Small
J. V.
(
2008
).
Building the actin cytoskeleton: filopodia contribute to the construction of contractile bundles in the lamella
.
J. Cell Biol.
180
,
1234
-
1244
.
Oser
M.
,
Condeelis
J.
(
2009
).
The cofilin activity cycle in lamellipodia and invadopodia
.
J. Cell. Biochem.
108
,
1252
-
1262
.
Oser
M.
,
Yamaguchi
H.
,
Mader
C. C.
,
Bravo-Cordero
J. J.
,
Arias
M.
,
Chen
X.
,
Desmarais
V.
,
van Rheenen
J.
,
Koleske
A. J.
,
Condeelis
J.
(
2009
).
Cortactin regulates cofilin and N-WASp activities to control the stages of invadopodium assembly and maturation
.
J. Cell Biol.
186
,
571
-
587
.
Otto
J. J.
,
Kane
R. E.
,
Bryan
J.
(
1979
).
Formation of filopodia in coelomocytes: localization of fascin, a 58,000 dalton actin cross-linking protein
.
Cell
17
,
285
-
293
.
Paavilainen
V. O.
,
Oksanen
E.
,
Goldman
A.
,
Lappalainen
P.
(
2008
).
Structure of the actin-depolymerizing factor homology domain in complex with actin
.
J. Cell Biol.
182
,
51
-
59
.
Pavlov
D.
,
Muhlrad
A.
,
Cooper
J.
,
Wear
M.
,
Reisler
E.
(
2007
).
Actin filament severing by cofilin
.
J. Mol. Biol.
365
,
1350
-
1358
.
Pollard
T. D.
,
Cooper
J. A.
(
2009
).
Actin, a central player in cell shape and movement
.
Science
326
,
1208
-
1212
.
Prochniewicz
E.
,
Janson
N.
,
Thomas
D. D.
,
De la Cruz
E. M.
(
2005
).
Cofilin increases the torsional flexibility and dynamics of actin filaments
.
J. Mol. Biol.
353
,
990
-
1000
.
Ressad
F.
,
Didry
D.
,
Xia
G. X.
,
Hong
Y.
,
Chua
N. H.
,
Pantaloni
D.
,
Carlier
M. F.
(
1998
).
Kinetic analysis of the interaction of actin-depolymerizing factor (ADF)/cofilin with G- and F-actins. Comparison of plant and human ADFs and effect of phosphorylation
.
J. Biol. Chem.
273
,
20894
-
20902
.
Romero
S.
,
Le Clainche
C.
,
Didry
D.
,
Egile
C.
,
Pantaloni
D.
,
Carlier
M. F.
(
2004
).
Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis
.
Cell
119
,
419
-
429
.
Romero
S.
,
Didry
D.
,
Larquet
E.
,
Boisset
N.
,
Pantaloni
D.
,
Carlier
M. F.
(
2007
).
How ATP hydrolysis controls filament assembly from profilin-actin: implication for formin processivity
.
J. Biol. Chem.
282
,
8435
-
8445
.
Schirenbeck
A.
,
Arasada
R.
,
Bretschneider
T.
,
Stradal
T. E.
,
Schleicher
M.
,
Faix
J.
(
2006
).
The bundling activity of vasodilator-stimulated phosphoprotein is required for filopodium formation
.
Proc. Natl. Acad. Sci. USA
103
,
7694
-
7699
.
Schmoller
K. M.
,
Semmrich
C.
,
Bausch
A. R.
(
2011
).
Slow down of actin depolymerization by cross-linking molecules
.
J. Struct. Biol.
173
,
350
-
357
.
Sheetz
M. P.
,
Wayne
D. B.
,
Pearlman
A. L.
(
1992
).
Extension of filopodia by motor-dependent actin assembly
.
Cell Motil. Cytoskeleton
22
,
160
-
169
.
Spudich
J. A.
,
Watt
S.
(
1971
).
The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin-troponin complex with actin and the proteolytic fragments of myosin
.
J. Biol. Chem.
246
,
4866
-
4871
.
Suarez
C.
,
Roland
J.
,
Boujemaa-Paterski
R.
,
Kang
H.
,
McCullough
B. R.
,
Reymann
A. C.
,
Guérin
C.
,
Martiel
J. L.
,
De La Cruz
E. M.
,
Blanchoin
L.
(
2011
).
Cofilin tunes the nucleotide state of actin filaments and severs at bare and decorated segment boundaries
.
Curr. Biol.
21
,
862
-
868
.
Svitkina
T. M.
,
Borisy
G. G.
(
1999
).
Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia
.
J. Cell Biol.
145
,
1009
-
1026
.
Urban
E.
,
Jacob
S.
,
Nemethova
M.
,
Resch
G. P.
,
Small
J. V.
(
2010
).
Electron tomography reveals unbranched networks of actin filaments in lamellipodia
.
Nat. Cell Biol.
12
,
429
-
435
.
Vignjevic
D.
,
Kojima
S.
,
Aratyn
Y.
,
Danciu
O.
,
Svitkina
T.
,
Borisy
G. G.
(
2006
).
Role of fascin in filopodial protrusion
.
J. Cell Biol.
174
,
863
-
875
.
Yamaguchi
H.
,
Lorenz
M.
,
Kempiak
S.
,
Sarmiento
C.
,
Coniglio
S.
,
Symons
M.
,
Segall
J.
,
Eddy
R.
,
Miki
H.
,
Takenawa
T.
,
Condeelis
J.
(
2005
).
Molecular mechanisms of invadopodium formation: the role of the N-WASP-Arp2/3 complex pathway and cofilin
.
J. Cell Biol.
168
,
441
-
452
.

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