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Vol. 18, Issue 10, 3928-3940, October 2007
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*Department of Cell and Molecular Biology, Feinberg School of Medicine, Northwestern University, Chicago, IL 60611; and Marine Biological Laboratory, Woods Hole, MA 02543
Submitted April 16, 2007;
Revised June 11, 2007;
Accepted July 19, 2007
Monitoring Editor: Josephine Adams
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Davenport et al., 1993; Dent and Gertler, 2003; Koleske, 2003). Filopodial turning, elongation, and retraction are key events involved in this process and require filopodia to be flexible enough to wave about yet rigid enough to protrude many microns past the cell surface. To identify the physical and molecular properties which underlie the reorganization of filopodia we must understand how actin filaments are cross-linked and how those cross-links are subsequently taken apart. Because fascin is the primary actin cross-linker in filopodia and is essential for filopodia formation (Vignjevic et al., 2006), we investigated features important to its function and dynamics. Specifically, we focused on questions regarding the initiation and elongation stages of filopodia formation as well as their maintenance and retraction: What is the origin of fascin in filopodia? How does fascin get delivered to tips of growing filopodia? How is it organized within actin bundles? How does that organization change over time, and what implications does this reorganization of filaments have on filopodia dynamics and stiffness?
Filopodia are thought to form by at least two mechanisms, one mechanism that is based on the reorganization of the dendritic network in a process that involves actin filament elongation and convergence at the barbed ends (Svitkina et al., 2003) and a formin-mediated process, which involves the formation of actin bundles in an unbranched network (Kovar and Pollard, 2004; Zigmond, 2004; Pellegrin and Mellor, 2005). Under the former mechanism, merged filaments elongate by polymerization of actin to form precursors to filopodia, so called lambda precursors. Actin filaments that come in proximity to one another are cross-linked into unipolar parallel bundles to form mature filopodia (Vignjevic et al., 2006). However, there is no established molecular mechanism that explains how fascin molecules are recruited to filopodia. One possibility is a higher affinity for actin filaments in filopodia than elsewhere in the cell. Activation by dephosphorylation is one way that has been reported to achieve this higher affinity (Ono et al., 1997; Yamakita et al., 1996; Adams et al., 1999; Vignjevic et al., 2006).
Cross-linking of actin filaments carries implications both for the elongation of filopodia and for their response to mechanical stress. Cross-linking is presumed essential for filopodial protrusion, because single actin filaments are insufficiently stiff to resist compressive forces (Mogilner and Rubinstein, 2005). Experimental support for this conclusion comes from observation of fascin-depleted filopodia that are typically bent and buckled under the cell membrane (Brieher et al., 2004; Vignjevic et al., 2006). It has been established that elongation of filopodia is mediated by actin assembly at their tips (Mallavarapu and Mitchison, 1999). Presumably, a supply of fascin is also required to cross-link the newly polymerized actin. Otherwise, the nascent actin filaments would not be sufficiently rigid to support protrusion of the filopodium. To reach the growing (barbed) end of nascent actin filaments, fascin must be translocated from the cell body into the filopodium and out to the tip. However, when filopodia reach long lengths or elongate rapidly, diffusion, alone, may not be a sufficient mechanism for protein transport. What other mechanisms does the cell invoke to overcome the limitations of simple diffusion?
If filaments are physically attached to each other by cross-linking, how can they reorganize to relieve intra- and extracellular stresses without compromising structural integrity? Others have shown that dynamic interactions exist between 
-actinin and actin filaments in stress fibers and that the dissociation rate of the cross-linker governs the mechanical properties of these bundles (Sato et al., 1987; Xu et al., 1998, 2000). We and others recently reported that fascin rapidly exchanges between filopodial actin filaments (Nakagawa et al., 2006; Vignjevic et al., 2006). The time-scale for fascin dissociation may provide clues as to how filopodia respond to mechanical stress that may have implications for the relationship of such response to cell morphology.
The purpose of this work is to advance our understanding of filopodia formation, which currently lacks detailed information regarding the fascin–actin interaction and how that interaction influences the molecular organization, remodeling, and mechanical properties of filopodia during its dynamic life cycle. Therefore, we undertook a quantitative, biophysical, and kinetic analysis of fascin dynamics. We quantified kinetic rates for fascin turnover in filopodia and we determined whether these rates are regulated or whether they are intrinsic to the cross-linker. We probed the role of phosphorylation cycles in regulating the active pool of fascin available for actin cross-linking by using phosphomimetic fascin mutants. Finally, we examined the limitations of fascin transport by diffusion and we developed a quantitative, computational model to analyze mechanisms for the delivery of cross-links to filopodial tips.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Tetramethylrhodamine (TMR)-actin was prepared by labeling F-actin with 5- (and 6)-carboxytetramethylrhodamine succinimidyl ester (NHSR) (Invitrogen, Carlsbad, CA) as described in Isambert et al. (1995) and stored at –80°C. Biotinylated actin was prepared with biocytin maleimide (Invitrogen) by the method of Rock et al. (2000). Before use, labeled G-actin was recycled by polymerization for 2 h on ice in the presence of 50 mM KCl, 2 mM MgCl, and 1 mM ATP, sedimentation at 100,000 x g for 1.5 h at 4°C, resuspension in cold G-buffer (2 mM Tris-HCl, 0.2 mM CaCl, 0.2 mM ATP, and 0.5 mM dithiothreitol [DTT]) to a final concentration of 2 mg/ml, and dialysis was conducted overnight against G-buffer using microdialysis buttons (Pierce Chemical, Rockford, IL).
Recombinant human fascin was prepared by a modification of the method of Ono et al. (1997). Escherichia coli carrying the plasmid was grown at 37°C until the A600 reached 0.6. Protein expression was induced by adding 0.1 mM isopropyl 
-D-thiogalactoside at 20°C for 4 h. Cells were harvested by centrifugation and extracted with B-PER in phosphate buffer (Pierce Chemical) plus 1 mM phenylmethylsulfonyl fluoride (PMSF) and 1 mM DTT. The lysate was centrifuged at 20,000 x g for 20 min, and the supernatant was mixed for 1 h at room temperature (RT) with 2 ml of glutathione-Sepharose 4B (GE Healthcare, Little Chalfont, Buckinghamshire, United Kingdom) equilibrated with phosphate-buffered saline (PBS) plus 1 mM DTT. The glutathione-Sepharose was poured into a column and washed with 20 ml of PBS plus 1 mM DTT. Then, 80 µl of thrombin (GE Healthcare) was added, and digestion was allowed to proceed overnight at 4°C. Flowthrough fractions were collected in 2 mM PMSF and concentrated by Centricon 10 (Millipore, Billerica, MA). Fascin was labeled with AlexaFluor 488 carboxylic acid, and succinimidyl ester (Alexa 488-NHS; Invitrogen). The pH of the fascin solution was raised to 8.3 by the addition of 1/20 volume of 0.1 M NaHCO3, pH 9.3. A 10-fold molar excess of Alexa 488-NHS was added from a stock solution of 25 mg/ml in dimethyl sulfoxide and reacted for 1 h at RT. Free dye was separated from labeled fascin on an Excellulose desalting column (Pierce Chemical).
In Vitro Bundling Assay
Coverslips (22 x 22 mm) were cleaned with 70% ethanol and blown dry with air (same applies to microscope slides). Coverslips were coated with nitrocellulose (1% collodion in amyl acetate (Electron Microscopy Sciences, Hatfield, PA) and allowed to air dry for 1 h. Chambers were created by placing two strips of double-sided tape onto microscope slides, 0.5 inch apart, and placing coverslip (nitrocellulose coating face down) on top of taped slide. Neutravidin mix (1 µl of 5 mg/ml Neutravidin in 50 mM PBS, pH 8.0, to 20 µl of actin polymerization buffer [10 mM imidazole, pH 7.0, 50 mM KCl, 1 mM MgCl2, and 1 mM EGTA]) was infused into perfusion chamber and allowed to incubate at RT for 1 min. Chamber was washed two times with 1 mg/ml bovine serum albumin in actin polymerization buffer to block any nonspecific binding. The bundle mixture (1 µM TMR-actin, 3 µM unlabeled actin, and 1 µM biotinylated-actin) was infused into chamber. Chamber inlet and outlet was sealed with petroleum jelly:lanolin:paraffin (1:1:1).
The bundle mixture was added to an Eppendorf tube and allowed to polymerize in actin polymerization buffer. To bundle actin filaments, 3 µM unlabeled fascin and 1.5 µM Alexa488-fascin were added to the test tube and allowed to incubate at RT for 1 h. The following were used to inhibit photobleaching: 4.5 mg/ml glucose, 0.86 mg/ml glucose oxidase (G-2133; Sigma-Aldrich, St. Louis, MO), and 0.14 mg/ml catalase (C-3155; Sigma-Aldrich) added to bundle mixture before imaging.
Cell Culture
B16F1 mouse melanoma and Neuro2a mouse neuroblastoma lines were provided by Drs. C. Ballestrem (Weizmann Institute of Science, Rehovot, Israel) and A. Ferreira (Northwestern University, Chicago, IL), respectively, and they were maintained in DMEM supplemented with 10% fetal bovine serum (FBS) at 37°C. Cells were plated onto coverslips, which were coated with 20 µg/ml mouse laminin (Invitrogen) for 1 h and thereafter incubated with 1% heat-inactivated bovine serum albumin in phosphate buffered saline for 20 min. pEGFP–fascin construct was provided by Dr. J. Adams (Cleveland Clinic Foundation, Cleveland, OH). QuikChangeII site-directed mutagenesis kit (Stratagene, La Jolla, CA) was used to create point mutations. Transfection of plasmids in B16 cells was carried out by FuGENE6 (Roche Diagnostics, Indianapolis, IN) and TransIT-Neural (Mirus, Madison, WI) for Neuro2a cells per the manufacturers' instructions. For live cell imaging, cells were transferred into L-15 medium (Invitrogen) supplemented with 10% FBS.
Quantitative Immunoblotting
B16 cells were lysed in buffer containing 10 mM Tris, pH 7.5, 150 mM NaCl, 1% Triton X-100, 10% glycerol, and protease inhibitor tablet (Roche Diagnostics). Lysate and purified fascin protein were loaded onto an SDS-polyacrylamide gel electrophoresis (PAGE) (4–20% polyacrylamide) and immunoblotted with anti-fascin primary antibody (DakoCytomation, Carpinteria, CA) and secondary mouse antibody (Sigma-Aldrich) using standard protocols. The amount of fascin in cell lysates was evaluated by comparing the intensity of the bands of each sample with the pure fascin standard by densitometry using NIH ImageJ software (http://rsb.info.nih.gov/ij/).
Microscopy
FRAP and Fluorescence Loss in Photobleaching (FLIP).
FRAP and FLIP experiments were performed on a Zeiss LSM 510 Meta confocal microscope (Carl Zeiss, Thornwood, NY) with a 100x/1.3 numerical aperture planapochromat oil objective. TMR-actin was excited with the 543-nm line of a HeNe laser (1 mW), whereas green fluorescent protein (GFP)- and Alexa488-tagged proteins were excited with the 488-nm line of an argon laser (25 mW). We photobleached fluorescence using a rectangular region of 1 µm2 area for 0.7–1.7 s by using 100% laser power for photobleaching, and we recorded images at attenuated laser settings.
FRAP data were analyzed as follows. The average intensity in the bleached zone was measured over the time course of the experiment. Before quantifying the fluorescence recovery, the measurements were corrected for photobleaching during image acquisition by a bleaching correction factor. This factor was determined by curve-fitting the fluorescence decay of a filopodium located far from the site of bleaching during a FRAP experiment (nonbleached bundle in the in vitro experiments) with a single exponential function, exp(–kdecayt), to extract a decay constant, kdecay. We then multiplied the intensity in the bleached zone by the correction factor, exp(–kdecayt), for each time point, t, in the experiment to obtain a normalized recovery profile. We curve fit the in vivo FRAP profile with a double exponential function, I(t) = If – 0.75 x exp(–kfastt) – 0.25 x exp(–kslowt), and single exponential function, I(t) = If + (Io – If)exp(–kfastt), for the in vitro experiment, because fluorescence recovered rapidly to prebleach values, where If is the intensity of the bleached zone at the final time of the experiment relative to the prebleach state, Io is the intensity immediately postbleach relative to the prebleach state, kfast is the constant reflecting the recovery to quasi-steady state when fascin cross-links have redistributed evenly within the filopodium, and kslow is the constant reflecting the recovery to prebleach intensity. The half time for dissociation was calculated as t1/2 = ln2/kfast, with average values along with their 95% confidence interval reported.
FLIP data were analyzed by measuring the loss in fluorescence near the filopodial tip. The intensity loss was multiplied by a photobleaching correction factor, which was determined in the same manner as above. Once normalized, the decay in fluorescence was fit with a single exponential function, I(t) = Io exp(–kofft), where Io is the initial intensity relative to the prebleach state and koff is the dissociation rate constant.
Epifluorescence Live Cell Imaging: Determination of Percentage of Fascin Localizing in Filopodia. B16 mouse melanoma cells expressing GFP-fascin were visualized by fluorescence microscopy using Nikon Diaphot CH-300 (Nikon, Tokyo, Japan), equipped with a 63x objective, at 300-ms exposure. Cells were kept on the microscope stage at 37°C during observation. Images were analyzed using MetaMorph imaging software (Molecular Devices, Sunnyvale, CA). MetaMorph line function was used to trace and measure the whole-cell perimeter.
Confocal Microscopy: Determination of Percentage of Fascin Bound in Filopodia. B16 mouse melanoma cells expressing GFP-fascin were imaged using the LSM510 Meta confocal microscope and kept at 37°C with consistent settings throughout imaging and analyzed using the LSM510 Meta image analyzer software (Carl Zeiss).
Determination of the Operating Fascin:actin Ratio in Filopodia
Quantitative immunoblotting was used to determine the concentration of cellular fascin protein to be 100–300 nM in B16 cells (350–500 nM in N2a cells). That value was converted to number of fascin molecules per cell by dividing the amount of fascin protein by molecular weight of fascin (55 kDa) and Avogrado's number, yielding 1.0–2.5 x 105 molecules per B16 cell (3.0–4.6 x 105 per N2a cell). Using fluorescence microscopy, we determined the percent of total fascin molecules in a cell which localize to filopodia, 11% in B16 (29% in N2a). To calculate the number of fascin molecules in one filopodium, we divided by the average number of filopodia per cell (number of filopodia in B16 cells = 30, n = 88; number of filopodia in N2a cells = 60, n = 30), or
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) and N2a filopodia were 8 µm (current study). By assuming that filopodia contain 20 actin filaments (Lewis and Bridgman, 1992; Mogilner and Rubinstein, 2005) and using the accepted incremental extension of an actin monomer of 2.7 nm, which gives 370 actin monomers per µm of filament, we found that the filopodia contain:
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) was performed in melanoma cells. To extend those studies to a neuronal cell type and to evaluate the conserved features of fascin exchange, we determined the dynamics of fascin in Neuroblastoma 2a cells, a cell line that generates numerous filopodia. We compared the distribution and turnover of fascin in filopodia by using fluorescence microscopy and FRAP analysis. We found that fascin localizes throughout the entire length of filopodia in these cells (Figure 1A), just as in B16F1 mouse melanoma cells (Vignjevic et al., 2006). In addition, FRAP experiments revealed rapid recovery of GFP-fascin in filopodia, on the order of 10 s (Figure 1B), and this behavior did not require the turnover of actin, which moved with retrograde flow (Figure 1C). The time scale for fascin exchange was consistent between phases of filopodial protrusion, retraction, and stationary periods. This result is consistent with findings in other cell types (Nakagawa et al., 2006; Vignjevic et al., 2006), which indicates that the dynamic behavior is a general phenomenon. The conserved behavior implies that the localization and dynamics of fascin may be critical to filopodia remodeling and intimately connected to their biological function.
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What is the significance of recovery that is uniform but to less than prebleach values? One possibility is that a fraction of the bleached molecules have been rendered immobile. Another possibility is the existence of a second, slower mode of recovery. Extending image acquisition to >100 s showed essentially full restoration of prebleach fluorescence (Figure 1E). Because we could infer that the initial rapid recovery was predominantly due to intrafilopodial exchange, the secondary slow recovery process suggests two additional conclusions: first, that essentially all the fascin has remained mobile, and, second, that fascin in filopodial cross-links can exchange with the cytoplasmic pool of fascin. FRAP experiments along the entire length of filopodia containing GFP-fascin revealed recovery times that matched those of the secondary recovery phase, confirming these conclusions (Supplemental Figure 1).
FRAP Data Reveal Kinetic Off-Rate of Fascin in Filopodia
What is the kinetic significance of the fast fascin recovery seen in the FRAP data? Is this recovery a measure of fascin association, dissociation, or a mixture of both events? Theoretical considerations indicate that in a simple binding reaction (A + B 
AB), recovery is exponential (1 – exp(–kofft)) and that the exponent koff indicates the rate constant of dissociation from the complex (Sprague et al., 2004). On this basis, we calculate an off rate of koff = 0.12 s–1.
We sought to confirm this value by independent FLIP experiments, which provided a more direct measure of the fascin dissociation rate constant in filopodia. Here, we repeatedly bleached a proximal domain of a filopodium containing GFP-fascin, while monitoring the loss of fluorescence over time in a distal domain (Figure 2A). Because the bleaching light was confined to the proximal domain, any loss of fluorescence in the distal domain was due to fascin molecules that dissociated from actin filaments in that domain and diffused into the proximal domain. If we assume that a fascin molecule typically diffused out of the distal zone before rebinding, the diminishing brightness of the distal domain was a direct indicator of the rate of fascin dissociation. The decay of fluorescence in the FLIP experiment was plotted versus time (Figure 2B) and curve-fit using a single exponential function, exp(–kofft), revealing an off rate, koff = 0.1 s–1 (n = 19), corresponding to a half-time for fascin dissociation of 
6 s. As expected, the half-time for dissociation matches the half time of recovery from FRAP studies, shown by the complementary profiles of the curves plotted in Figure 2B, reinforcing the conclusion that fascin dissociates from actin filaments with a koff = 0.12 s–1.
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), thereby excluding any proteins that potentially regulate fascin exchange. If the recovery behavior of fascin in purified bundles matched that in filopodia, then it would be evidence that fascin turnover is an intrinsic property of the cross-linker. We tethered in vitro bundles to coverslips via a biotin–avidin interaction in a sealed perfusion chamber so that fluorescence recovery could be monitored over time (Figure 3A). FRAP experiments revealed rapid fluorescence recovery of Alexa488-fascin (Figure 3B, top). In contrast, fluorescence recovery of TMR-actin was negligible (Figures 3B, bottom, and C), confirming our in vivo result that fascin undergoes cycles of dissociation and association within actin bundles. The recovery data were normalized for global bleaching and fit with a single exponential function as shown in Figure 3C. The half time of recovery for fascin in vitro is t1/2 = 6 ± 3 s (n = 28), nearly identical to that found in filopodia (Figure 3D). Because the time scale for fascin exchange was recapitulated in vitro, we conclude that fascin exchange is an intrinsic property of the fascin–actin interaction and that it does not require assistance by an auxiliary molecule(s) for either binding or release from filopodial actin.
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20 s, whereas full filopodial recovery required a slower component of exchange between the filopodium and cell body. The rapid full recovery in vitro was expected, as the bundles were surrounded by a large unbound pool of fascin that could undergo three-dimensional diffusion over shorter distances to unoccupied binding sites. In contrast, full filopodial recovery required long-distance exchange with the cell body and diffusion through a more crowded space.
Because such an intrinsic property precludes a multistep association reaction (e.g., with a catalyst), we can use the value of a previously measured affinity constant (KA = 6.7 µM–1, Yamakita et al., 1996) and our measured koff to calculate the kinetic on-rate, kon, of the fascin–actin association reaction as follows: kon = KA· koff = 0.8 µM–1 s–1.
Fascin-Mediated Actin Bundling in Filopodia Requires That Fascin Be in the Nonphosphorylated State
Several studies have shown that the formation and lifetime of filopodia seem dependent upon the state of fascin phosphorylation; when activated by dephosphorylation, fascin yielded numerous, long-lived filopodia, whereas phospho-fascin yielded sparse, short-lived filopodia (Yamakita et al., 1996; Adams et al., 1999; Anilkumar et al., 2003; Vignjevic et al., 2006). These observations suggest that phosphorylation limits the availability of fascin for actin bundling in filopodia. However, phosphorylated fascin is not excluded from filopodia (Vignjevic et al., 2006); therefore, its contribution to actin bundling needs to be tested. Here, we quantified and compared the preference for localization in filopodia and the turnover rates of fascin phospho-mimetic mutants to determine the extent of binding of phosphorylated fascin in filopodia.
To directly test whether the incorporation of fascin in filopodia was dependent on the phosphorylation state, we analyzed the fluorescence distribution of soluble GFP, GFP-tagged wild-type fascin, and GFP-tagged mutant fascin proteins that mimic the inactive phosphorylated (S39E) and active nonphosphorylated (S39A) states (Figure 4A). Proteins were expressed in N2a cells (on a background of endogenous levels), because these cells generate many filopodia that are easy to study. We used confocal microscopy to obtain images of consistent optical depth so that measured intensities would indicate relative GFP concentrations, and we plotted the ratio of filopodial-to-lamellipodial fluorescence for each protein in Figure 4B. As a control for uniform optical depth, we first measured this ratio for GFP-expressing cells, and we found that Ifil/Ilam = 0.9 ± 0.1 1 (n = 26), as expected for a uniformly soluble marker. Also as expected, wild-type GFP-fascin showed significantly greater preference for filopodia (Ifil/Ilam = 7.4 ± 1.0; n = 27) than did diffuse GFP. Although both mutants localized along filopodial shafts, the active S39A mutant showed strong preference for filopodia (Ifil/Ilam = 10.0 ± 2.9; n = 27), whereas filopodia containing inactive S39E exhibited only marginal localization (Ifil/Ilam = 1.3 ± 0.1; n = 27) and a uniform diffuse fluorescence in the cell body. The comparison of intensity distribution among fascin mutants implies that fascin incorporation in filopodia is strongly increased by fascin dephosphorylation.
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). These results suggest that the bound fascin cross-links in filopodia exist predominantly in the dephosphorylated state and that this feature is conserved among neuronal and nonneuronal cells. In contrast, the recovery profile for the GFP-tagged S39E mutant (Figure 4D, left) closely resembled that of soluble GFP (Figure 4D, right), with t1/2 = 1.7 ± 0.6 s (n = 13) and t1/2 = 0.5 ± 0.2 s (n = 10), respectively (Figure 4E). Instead of revealing a distinct bleached zone, as in the wild-type and S39A FRAP experiments, the bleached zone and the region distal to it seemed to be generally nonfluorescent after photobleaching despite bleaching a zone of identical size (1-µm2 area). We infer from the similar mobilities of GFP and the S39E mutant that neither protein appreciably bind actin. Instead, both GFP and S39E likely diffused into the bleached zone during the bleaching interval before redistributing into the distal domain of the filopodium, leaving the bleached zone and the region distal to it nonfluorescent.
Fascin Undergoes Free Diffusion in Filopodia with a Moderate Diffusion Coefficient, D = 6 µm2/s
The tight packing of actin filaments and closely apposed plasma membrane of filopodia suggest an environment of limited diffusive mobility. We sought a measure of the diffusion coefficient of fascin in filopodia to understand the relative difficulty of movement compared with that in the cell body. Using experimentally derived images of FRAP as inputs of filopodia cross-sections and initial and final protein concentrations, we simulated the recovery process of both GFP and GFP-S39E by using a mathematical model of one-dimensional diffusion (Appendix A). Typical experimental recovery profiles and their simulation results are plotted versus time in Figure 5A. By matching the recovery half-times between experiment and simulation, we determined values of mean diffusion (D) coefficient for GFP and GFP-S39E of 7.5 µm2/s and 5.1 µm2/s, respectively. Given the purely diffusive motion of soluble GFP, the Stokes–Einstein relationship predicts a diminished diffusion coefficient with increasing molecular weight: D 
M–1/3. We fit this relationship through the experimentally determined diffusion coefficient of GFP, and we plotted the experimental value for GFP-S39E on the same plot (Figure 5B). The match between the experimentally measured diffusion coefficient for GFP-S39E and the predicted value based on molecular weight confirms that the phosphomimetic protein is not measurably slowed by actin binding or small network pore size in filopodia. Had the recovery process for GFP-S39E been significantly slowed by kinetics, the measured D value would have fallen below the curve.
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), reflecting the increased rate of collisions with immobile proteins, the more tortuous paths within the tight filopodial network, or both.
Incorporation of Fascin Cross-links in Filopodia
Individual actin filaments cannot impart sufficient stiffness to filopodia extended beyond the leading edge, but bundling by fascin allows for protrusion and maintenance of filopodia (Brieher et al., 2004; Mogilner and Rubinstein, 2005; Vignjevic et al., 2006). It is not known, however, what the fascin-to-actin ratio is in filopodia. Additionally, although our FRAP data reveal the existence of bound and free populations of fascin in filopodia, we do not know the fraction bound. We designed two approaches to calculate the relative fascin to actin concentration and solved for the ratio of bound to free fascin. Together, these relationships provide an estimate for the number of bound fascin molecules per actin in filopodia.
Filopodia Contain One Fascin Cross-link per 25–60 Actin Monomers
To solve for the stoichiometry of fascin to actin in filopodia, we developed a two-part technique that involves quantitative immunoblotting and fluorescence ratiometry based on previously established methods (Zhai and Borisy, 1994; Wu and Pollard, 2005). Immunoblotting was used to calculate the total number of fascin molecules in cells, and fluorescence microscopy was performed to determine the proportion of cellular fascin molecules that localize to filopodia. The measurements were combined to calculate the number of fascin cross-links per actin residing in filopodia.
To determine the concentration of fascin in a cell, nontransfected cells were plated in a dish and counted by phase microscopy. Lysates of the counted cell population along with known amounts of pure fascin protein were loaded onto an SDS-PAGE and incubated with fascin antibody. The blot was visualized by chemiluminescence and shown in Figure 6A and Supplemental Figure 2. The intensity of the bands corresponding to amount of fascin in cell lysate and pure protein were quantified using NIH image software. We determined that B16 and N2a cells contain roughly 100–500 nM (n = 20).
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10–30% (n = 118). Together, quantitative immunoblotting and fluorescence fractionation revealed that average filopodia contain one fascin cross-link per 25–60 actin monomers (calculation shown in Materials and Methods).
Most Fascin Molecules in Filopodia Are Bound at Any Instant
The stoichiometry of fascin to actin in filopodia determined above includes the total population of fascin molecules in filopodia; both bound and unbound fractions. Here, we used a fluorescence ratiometric technique to determine the proportion of fascin molecules that are bound. We examined the difference between intensity in filopodia and cytoplasm in confocal images of cells expressing GFP-fascin at varying expression levels. Confocal microscopy serves as an advantage to wide-field microscopy in that an image of the same optical depth is acquired. Therefore, intensity levels between different cellular structures, namely, filopodia and cytoplasm, can be directly compared. Figure 7A shows a representative confocal image of a B16 cell expressing GFP-fascin. Based on the assumption that the cytoplasmic pool in the cell body is unbound and that this concentration is uniform throughout the cell and filopodia, the per-pixel fluorescence intensity in the cell body (dashed circular region in Figure 7B) was taken as proportional to free fascin concentrations, whereas the fluorescence intensity in the filopodium (solid black outline in Figure 7B) was taken as proportional to the sum of bound and free fascin. The assumption that fascin is predominantly soluble in the cell body is supported by FRAP experiments in which rapid diffusion precluded any discernible bleached zone within lamellipodia or the cell body (data not shown). Therefore, dividing the fluorescence intensity in the cell body by the filopodial intensity yields the proportion of unbound fascin population in the filopodium. Subtracting this value from 1 gives the proportion of bound fascin molecules in filopodia. We plotted the percentage of bound fascin molecules in filopodia against average intensity values in the cytoplasm of cells at varying expression levels (Figure 7C and Supplemental Figure 2C). To determine the percentage of bound fascin molecules in wild-type filopodia, we curve fit our data with a second order polynomial and extrapolated to zero cell intensity. The y-intercept of the curve revealed that, on average, 97% (B16 cells: ncells = 51, nfil = 615; N2a cells: ncells = 26, nfil = 353) of endogenous fascin molecules are bound in filopodia, with a range of 94–98%.
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1.2 µm. In filopodia significantly longer than this distance and with irreversible cross-links (i.e., koff = 0), diffusing fascin would permanently fill binding spots nearest the base first, and an elongation rate that exceeds fascin supply would leave the tips very low in fascin (Figure 8A). This rapid local transport but slow long-distance diffusion is the same phenomenon observed in the fast and slow recovery components in Figure 1. Diffusion with dynamic cross-linking (koff > 0) might permit a sufficient supply rate to the tip region at longer lengths, by allowing dissociation of bound fascin and continued diffusion and redistribution along the length of the filopodium.
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As expected, the advantage of fascin dissociation increased with filopodial length at typical rates of elongation. With irreversible cross-links, the profile of bound fascin concentration fell sharply toward the tip of 3-µm-long filopodia (Figure 8B, black curve). With dynamic cross-linking, the concentration of occupied sites near the base decreased due to a lower equilibrium (i.e., maximum) value, but increased by a factor of 1.5 near the tip (Figure 8B, gray curve). When filopodia continued to grow to 10 µm, a markedly sigmoidal concentration profile indicated low fascin transport to the tip when koff = 0 s–1, but a 100-fold increase in bound fascin concentration was observed with koff = 0.12 s–1. Thus, a process which involves diffusion and rapid, irreversible association can supply fascin only to short filopodia, but dynamic cross-linking is required to sustain longer filopodia.
Is there an optimal off-rate for delivery of fascin to filopodial tips? Figure 8C plots the concentrations of bound fascin [FS] near the tips of growing, 3-µm-long filopodia as a function of koff. Consistent with Figure 8B, tip concentrations initially increased with koff. At higher koff; however, concentrations decreased due to a diminishing equilibrium bound fascin concentration. The peaks in the curves represent an optimal value of koff for maximum bound fascin concentration near the tips of growing filopodia. Our measured off-rate compares well with the range of dissociation rates, 0.05–0.25 s–1, that are optimal for filopodia growing at rates of 1–4 µm/min.
Figure 8D plots the optimal koff as a function of length for several growth velocities. At growth rates of 2 µm/min and lengths of 3 µm, our measured value of koff = 0.12 s–1 provides the highest bound fascin concentration near the tip. At 3 µm/min filopodial growth rate, koff is optimized for short, 0.8 µm filopodia. At slow growth rates of 1 µm/min, bound fascin concentrations near the tip would be higher with a decreased koff at any length. Thus, the koff measured in this study seems to be a good compromise among lengths to generate the highest tip concentrations of bound fascin at typical growth rates.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Fascin Phosphorylation Acts as a Molecular Switch for Filopodia Formation
Cells have the capacity to form filopodia in response to a variety of signals that arise from environmental cues, particular substrates, or other cells. Such a capacity implies the existence of a regulatory mechanism. In this work, we tested phosphorylation/dephosphorylation cycles in regulating key events in filopodia. Because fascin phosphorylation has been reported to inhibit actin bundling (Yamakita et al., 1996; Ono et al., 1997; Adams et al., 1999), we suspected that the role of these cycles may be to regulate fascin exchange in filopodia and to limit concentrations of fascin available for actin cross-linking. Unexpectedly, we found that the time scale for fascin dissociation in reconstituted filopodial bundles mimicked those in filopodia, indicating that the dynamic exchange does not require an enzymatic reaction but that it is an intrinsic property of the fascin cross-linker. In contrast, photobleaching data on fascin phosphomutants revealed that phosphorylation alters the mobility of fascin in filopodia. Although the nonphosphorylated fascin mutant imitated wild-type kinetic reactions in filopodia, the phospho-fascin mutant underwent a predominantly diffusive process. The phosphomimetic mutant S39E contains one active binding site and one site that is inactivated by a single-site mutation. Although the nonmutated binding site is presumably free to associate with actin, it is likely that, without the cooperation of the other binding site, fascin does not undergo appreciable binding to filaments. Therefore, only the nonphosphorylated form seems to participate in filament cross-linking, whereas the purpose of fascin phosphorylation is likely to limit the size of that pool. On basis of our results, we propose that phosphorylation/dephosphorylation cycles serve as a molecular switch to turn fascin activity on or off, which in so doing determines whether filopodia form.
In summary, we propose a model for the formation and maintenance of filopodia (Figure 9). In our model, activation of fascin by dephosphorylation allows for high-affinity actin binding in filopodia. Actin cross-linking provides filopodia with the rigidity necessary for protrusion, whereas intrinsic fascin exchange between filaments allows for elongation and filopodial remodeling. Fascin inactivation by phosphorylation decreases the availability of fascin to bundle actin, which in turn inhibits filopodia formation.
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; Tilney et al., 2000; Volkman et al., 2001), and bound ratios of up to 1:4.6 have been measured in vitro (Bryan and Kane, 1978; Yamashiro-Matsumura and Matsumura, 1985). However, it seems unlikely that the geometry of filopodia would allow for this precise packing. The pitch of each actin helix of a free filament is imperfect for hexagonal packing, rotating 360° per 13.7 monomers, or only 26.3° (instead of 30°) per monomer. Because one bound fascin per 25–60 actin monomers equates to one cross-link every 6th to 15th monomer along a double helix, a torque aligning the filament for hexagonal packing would result in an cumulative angle strain of (30–26.3° per monomer) x (6–15 monomers) = 22–56° between successive cross-links on a filament. Electron micrographs of filopodia formed by convergent elongation in fact seem imperfectly packed (Svitkina et al., 2003), and Stokes and DeRosier (1991) have described bundles formed in vitro with one bound fascin per 33–66 actin monomers as plentiful but "poorly ordered," both suggesting a somewhat reduced binding site concentration. The lower range of our bound fascin-to-actin ratio (1:25) would translate to a binding site to actin ratio of 1:4.6 if either KA was as low as 2.3 µM–1 or the free fascin concentration, [F], was as low as 0.03 µM (an order of magnitude lower than our measured 0.1–0.5 µM). In summary, filopodia seem to represent semiordered bundles, reducing the possible fascin:actin (saturation) stoichiometry from a 1:4 theoretical value to 1:10–24. However, given their dynamic nature and limited soluble concentration, they only operate at ratios of 1:25–60 in vivo.
Fascin Dynamics Impart Rate-dependent Resistance to Filopodial Deformation
A key function of fascin as an actin cross-linker is its ability to stiffen filopodia. Individual actin filaments are insufficiently rigid to resist compressive forces (Janmey et al., 1991; Mogilner and Rubinstein, 2005), and filopodia in fascin-depleted cells are typically bent and buckle under the cell membrane (Vignjevic et al., 2006). Although the function of stiffening would apparently be best served by high-affinity fascin binding, this counteracts the need for filopodia to undergo necessary remodeling (e.g., fusion of neighboring filopodia, accommodation to obstacles, or cycling between growth and breakdown). The dynamic nature of the fascin cross-links allows for both requirements.
The filopodium as a whole undergoes two main loading regimes, with a different mechanical response to each. Slow-scale displacement of the filopodium occurs on the time scales of cell motility itself, on the order of 1 µm/min, and it may occur over small or large distances. A growing filopodium may be displaced as it encounters other cells, for example, or be displaced by lamellipodial retraction or the motion of adjacent cells. Displacement may also occur on rapid time scales due to Brownian (thermal) motion, under which a single, 10-µm-long actin filament bends considerably (persistence length, Isambert et al., 1995). We have shown that the half-time of fascin exchange in filopodia is 6 s, indicating a time scale of motion that divides two types of mechanical responses: displacement rates that build little strain in the cross-links over this time-scale result in little resistance to displacement from those cross-links, whereas rates that build up significant strain on this time scale encounter resistance due to the additional stiffness that fascin imparts to the bundle. This resistance rises with the filopodial fascin:actin ratio, which we estimated to be high, at 1:25–60. Although it is difficult to predict the rate at which strain builds in the cross-links of a displaced filopodium without detailed knowledge of the system geometry, it is likely that the slow displacements do not build appreciable resistance from fascin over the course of a single binding period, whereas the displacement due to thermal motion almost certainly does. The dynamics of fascin exchange therefore mediate a differential, visco-elastic-like response to the two common loads, allowing slow remodeling but resisting rapid fluctuations with additional stiffness (Figure 10).
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; Pollard et al., 1990). Like fascin-mediated actin bundles, the mechanical properties of such actin networks depend on the rate of displacement, where the stiffness can increase by an order of magnitude at fast displacements. One significant difference is that the rate of dissociation of -actinin is an order of magnitude faster than that of fascin (Xu et al., 2000). The variance in the time scales of protein exchange between the two bundlers yield different limits for the rate of deformation. Such limits may be critical for the ability of actin bundles to withstand the different types of stresses they encounter in their respective subcellular locations. It would be interesting to compare such properties between filopodia and other protrusive actin networks such as pseudopodia, microvilli, and stereocillia.
Fascin Turnover Is a Key Process in Rapid Filopodial Assembly
Filopodia are naturally subject to selective evolutionary pressures as dictated by their required sensory functions. Such pressures include the formation and maintenance of ordered parallel bundles; remodeling of filopodial structure at adequate rates under appropriate loads, with resistance to other loads; providing the necessary, perhaps nonuniform, stiffness along the filopodial length; and transport of cytoskeletal proteins to growing filopodial tips at rates compatible with elongation velocities. Such requirements must operate over filopodial lengths of 1–50 µm, diameters of only 0.1–0.5 µm (Mitchison and Cramer, 1996), and growth rates from a typical 2 µm/min (Mallavarapu and Mitchison, 1999; current study) to as fast as 25 µm/min (Sheetz et al., 1992). As our numerical analysis shows, filopodia growing at 2 µm/min seem to operate near a koff that delivers maximum fascin cross-links to the extending tip. Lower koff values allow less dissociation of bound fascin and continued diffusion toward the tip, whereas higher values further limit the equilibrium bound concentration throughout the length. Among the pressures affecting filopodial rigidity and morphology, the optimal value of koff to supply fascin to the tip at the typical growth rate seems to be of particular importance. Altogether, filopodia may have selected dynamic cross-links to form adequately stiff bundles at appropriate rates and allow for remodeling.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Yvonne S. Aratyn (y-aratyn{at}northwestern.edu)
Abbreviations used: FLIP, fluorescence loss in photobleaching; FRAP, fluorescence recovery after photobleaching; GFP, green fluorescent protein; RT, room temperature; TMR, tetramethylrhodamine; WT, wild type.
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REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Albrecht-Buehler, G. (1976). Filopodia of spreading 3T3 cells. Do they have a substrate-exploring function? J. Cell Biol 69, 275–286.[CrossRef]
Anilkumar, N., Parsons, M., Monk, R., Ng, T., and Adams, J. C. (2003). Interaction of fascin and protein kinase C: a novel intersection in cell adhesion and motility. EMBO J 22, 5390–5402.[CrossRef][Medline]
Brieher, W. M., Coughlin, M., and Mitchison, T. J. (2004). Fascin-mediated propulsion of Listeria monocytogenes independent of frequent nucleation by the Arp2/3 complex. J. Cell Biol 165, 233–242.
Bryan, J., and Kane, R. E. (1978). Separation and interaction of the major components of sea urchin actin gel. J. Mol. Biol 125, 207–224.[CrossRef][Medline]
Davenport, R. W., Dou, P., Rehder, V., and Kater, S. B. (1993). A sensory role for neuronal growth cone filopodia. Nature 361, 721–724.[CrossRef][Medline]
Dent, E. W., and Gertler, F. B. (2003). Cytoskeletal dynamics and transport in growth cone motility and axon guidance. Neuron 40, 209–227.[CrossRef][Medline]
Isambert, H., Venier, P., Maggs, A. C., Fattoum, A., Kassab, R., Pantaloni, D., and Carlier, M. F. (1995). Flexibility of actin filaments derived from thermal fluctuations. Effect of bound nucleotide, phalloidin, and muscle regulatory proteins. J. Biol. Chem 270, 11437–11444.
Janmey, P. A., Euteneuer, U., Traub, P., and Schliwa, M. (1991). Viscoelastic properties of vimentin compared with other filamentous biopolymer networks. J. Cell Biol 113, 155–160.
Koleske, A. J. (2003). Do filopodia enable the growth cone to find its way? Science STKE 2003, (183), pe20.
Kovar, D. R., and Pollard, T. D. (2004). Progressing actin: Formin as a processive elongation machine. Nat. Cell Biol 6, 1158–1159.[CrossRef][Medline]
Lewis, A. K., and Bridgman, P. C. (1992). Nerve growth cone lamellipodia contain two populations of actin filaments that differ in organization and polarity. J. Cell Biol 119, 1219–1243.
Mallavarapu, A., and Mitchison, T. (1999). Regulated actin cytoskeleton assembly at filopodium tips controls their extension and retraction. J. Cell Biol 146, 1097–1106.
Mitchison, T. J., and Cramer, L. P. (1996). Actin-based cell motility and cell locomotion. Cell 84, 371–379.[CrossRef][Medline]
Mogilner, A., and Rubinstein, B. (2005). The physics of filopodial protrusion. Biophys. J 89, 782–795.[CrossRef][Medline]
Nakagawa, H., Terasaki, A. G., Suzuki, H., Ohashi, K., and Miyamoto, S. (2006). Short-term retention of actin filament binding proteins on lamellipodial actin bundles. FEBS Lett 580, 3223–3228.[CrossRef][Medline]
Ono, S., Yamakita, Y., Yamashiro, S., Matsudaira, P. T., Gnarra, J. R., Obinata, T., and Matsumura, F. (1997). Identification of an actin binding region and a protein kinase C phosphorylation site on human fascin. J. Biol. Chem 272, 2527–2533.
Pollard, T. D., Satterwhite, L., Cisek, L., Corden, J., Sato, M., and Maupin, P. (1990). Actin and myosin biochemistry in relation to cytokinesis. Ann. NY Acad. Sci 582, 120–130.[Medline]
Pellegrin, S., and Mellor, H. (2005). The Rho family GTPase Rif induces filopodia through mDia2. Curr. Biol 15, 129–133.[CrossRef][Medline]
Rock, R. S., Rief, M., Mehta, A. D., and Spudich, J. A. (2000). In vitro assays of processive myosin motors. Methods 22, 373–381.[CrossRef][Medline]
Sheetz, M. P., Wayne, D. B., and Pearlman, A. L. (1992). Extension of filopodia by motor-dependent actin assembly. Cell Motil. Cytoskeleton 22, 160–169.[CrossRef][Medline]
Sato, M., Schwarz, W. H., and Pollard, T. D. (1987). Dependence of the mechanical properties of actin/-actinin gels on deformation rate. Nature 325, 828–830.[CrossRef][Medline]
Sprague, B. L., Pego, R. L., Stavreva, D. A., and McNally, J. G. (2004). Analysis of binding reactions by fluorescence recovery after photobleaching. Biophys. J 86, 3473–3495.[CrossRef][Medline]
Spudich, J., and Watt, S. (1971). The regulation of rabbit skeletal muscle contraction. J. Biol. Chem 246, 4866–4871.
Stokes, D. L., and DeRosier, D. J. (1991). Growth conditions control the size and order of actin bundles in vitro. Biophys. J 59, 456–465.[Medline]
Svitkina, T. M., Bulanova, E. A., Chaga, O. Y., Vignjevic, D. M., Kojima, S., Vasiliev, J. M., and Borisy, G. G. (2003). Mechanism of filopodia initiation by reorganization of a dendritic network. J. Cell Biol 160, 409–421.
Swaminathan, R., Hoang, C. P., and Verkman, A. S. (1997). Photobleaching recovery and anisotropy decay of green fluorescent protein GFP-S65T in solution and cells: cytoplasmic viscosity probed by green fluorescent protein translational and rotational diffusion. Biophys. J 72, 1900–1907.[Medline]
Tilney, L. G., and DeRosier, D. J. (1986). Actin filaments, stereocilia, and hair cells of the bird cochlea IV. How the actin filaments become organized in developing stereocilia and in the cuticular plat. Dev. Biol 116, 119–129.[CrossRef][Medline]
Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K., and Guild, G. M. (2000). Regulation of actin filament cross-linking and bundle shape in Drosophila bristles. J. Cell Biol 148, 87–100.
Vignjevic, D., Yarar, D., Welch, M. D., Peloquin, J., Svitkina, T., and Borisy, G. G. (2003). Formation of filopodia-like bundles in vitro from a dendritic network. J. Cell Biol 160, 951–962.
Vignjevic, D., Kojima, S. I., Aratyn, Y., Danciu, O., and Borisy, G. (2006). Role of fascin in filopodial protrusion. J. Cell Biol 174, 863–875.
Volkmann, N., DeRosier, D., Matsudaira, P., and Hanein, D. (2001). An atomic model of actin filaments cross-linked by fimbrin and its implications for bundle assembly and function. J. Cell Biol 153, 974–956.
Wu, J. Q., and Pollard, T. D. (2005). Counting cytokinesis proteins globally and locally in fission yeast. Science 310, 310–314.
Xu, J., Tseng, Y., and Wirtz, D. (2000). Strain hardening of actin filament networks; regulation by the dynamic cross-linking protein a-actinin. J. Biol. Chem 275, 35886–35892.
Xu, J, Wirtz, D., and Pollard, T. D. (1998). Dynamic cross-linking by a-actinin determines properties of actin filament networks. J. Biol. Chem 273, 9570–9576.
Yamakita, Y., Ono, S., Matsumura, F., and Yamashiro, S. (1996). Phosphorylation of human fascin inhibits its actin binding and bundling activities. J. Biol. Chem 271, 12632–12638.
Yamashiro-Matsumura, S., and Matsumura, F. (1985). Purification and characterization of an F-actin-bundling 55-kilodalton protein from HeLa cells. J. Biol. Chem 260, 5087–5097.
Zhai, Y., and Borisy, G. G. (1994). Quantitative determination of the proportion of microtubule polymer present during the mitosis-interphase transition. J. Cell Sci 107, 881–890.[Abstract]
Zigmond, S. H. (2004). Formin-induced nucleation of actin filaments. Curr. Opin. Cell Biol 16, 99–105.[CrossRef][Medline]
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