Research Article 1533

IntroductionActin filaments can assemble into diverse structures such aslamellipodia, filopodia or stress fibers. These structures in turnprovide protrusive and contractile forces during cell polarization,migration and division, and are central to many physiological andpathological processes such as differentiation, wound healingor malignant transformation. The driving force behind the oftenhighly dynamic behavior of cortical actin is a combination ofpolymerization, depolymerization and myosin-dependent motilityof actin filaments.

Polymerization of actin filaments relies on a variety of nucleationfactors. Among the different types of nucleators, formins constitutethe largest class. In addition to their nucleation activity, forminsalso have the ability to promote actin filament elongation. Thisdual function might explain why they have diverged so extensively(Chalkia et al., 2008). Formins support actin polymerization invitro at rates of up to 10 subunits/second at 1 M G-actin (Kovarand Pollard, 2004). By contrast, studies in a range of organismsindicate that formins facilitate in vivo rates of actin polymerizationof more than 550 subunits/second or 1.5 m/second (Higashida etal., 2004; Diez et al., 2005; Staiger et al., 2009). Importantly,although we understand formin biochemistry in great detail (Kovar,2006; Paul and Pollard, 2009), it has not been possible to link theelongation rate of a defined actin filament or bundle to the actionof a particular formin in vivo.

Myosin motors also contribute to actin remodeling by movingfilaments past one another or along fixed substrates. The generationand role of myosin-II-dependent contractile forces during musclecontraction (Yanagida, 2007) and cytokinesis (Wang, 2001; Wu etal., 2006; Clayton et al., 2010; Skau and Kovar, 2010) have beenintensely studied. Myosin II has also been implicated in retrogradeactin flow in budding yeast Saccharomyces cerevisiae (Huckaba et

al., 2006). Although there are several reports suggesting thatunconventional myosins could move actin filaments along the cellcortex, no direct evidence for such a function has yet been presented.Considering the ability of all myosins to move actin filaments in invitro gliding assays, and the association of many myosins with theplasma membrane (Zot and Pollard, 1993; Sousa and Cheney, 2005;McConnell and Tyska, 2007), this is quite remarkable.

The budding yeast S. cerevisiae has been an invaluable modelsystem for the study of molecular components and mechanismsregulating formin-dependent actin assembly (Evangelista et al., 2002;Sagot et al., 2002; Moseley and Goode, 2006). Actin cables in S.cerevisiae are linear filamentous structures generated by the twoformins, Bni1 and Bnr1. Bni1 localizes to the bud tip during polarizedgrowth and nucleates short, bud-localized cables. During cytokinesisBni1 relocates to the bud neck, where it supports assembly of cablesand, possibly, the acto-myosin ring (Evangelista et al., 2002; Butteryet al., 2007). Bnr1 is immobilized at the bud neck by interactionswith septins, and nucleates cables that project into the mother cell(Buttery et al., 2007). Polarized cables are used as tracks for myosin-V-driven vectorial transport, which is essential for cell growth andhomeostasis (Jansen et al., 1996; Pruyne et al., 1998).

Although actin cables are highly dynamic (Ayscough et al.,1997; Karpova et al., 1998), previous studies have focused onrelatively slow actin dynamics displayed by polarized cells (Yangand Pon, 2002; Huckaba et al., 2006). Dynamics of cables inunpolarized cells and during cell polarization remaineduncharacterized. Furthermore, the molecular mechanisms drivingcable dynamics remain unknown. Here, we use total internalreflection microscopy (TIRFM) to quantify and compare cabledynamics in unpolarized and polarized cells. Combiningquantitative image-analyses with genetic manipulations, we probethe molecular basis for the observed dynamics.

SummaryCell morphogenesis requires complex and rapid reorganization of the actin cytoskeleton. The budding yeast Saccharomyces cerevisiaeis an invaluable model system for studying molecular mechanisms driving actin dynamics. Actin cables in yeast are formin-generatedlinear actin arrays that serve as tracks for directed intracellular transport by type V myosins. Cables are constantly reorganizedthroughout the cell cycle but the molecular basis for such dynamics remains poorly understood. By combining total internal reflectionmicroscopy, quantitative image analyses and genetic manipulations we identify kinetically distinct subpopulations of cables that aredifferentially driven by formins and myosin. Bni1 drives elongation of randomly oriented actin cables in unpolarized cells, whereasboth formins Bnr1 and Bni1 mediate slower polymerization of cables in polarized cells. Type V myosin Myo2 surprisingly acts as amotor for translational cable motility along the cell cortex. During polarization, cells change from fast to slow cable dynamics throughspatio-temporal regulation of Bni1, Bnr1 and Myo2. In summary, we identify molecular mechanisms for the regulation of cabledynamics and suggest that fast actin reorganization is necessary for fidelity of cell polarization.

Key words: Actin dynamics, Cell polarity, Formin, MyoV

Accepted 8 December 2010Journal of Cell Science 124, 1533-1541 © 2011. Published by The Company of Biologists Ltddoi:10.1242/jcs.079038

Cortical actin dynamics driven by formins andmyosin VJerry H. Yu, Alvaro H. Crevenna, Mario Bettenbühl*, Tina Freisinger and Roland Wedlich-Söldner‡

AG Cellular Dynamics and Cell Patterning, Max Planck Institute of Biochemistry, Am Klopferspitz 18, 82152 Martinsried, Germany*Present address: Department of Mathematics, Potsdam University, Karl-Liebknecht-Strasse 24–25, 14476 Potsdam, Germany‡Author for correspondence (wedlich@biochem.mpg.de)

Jour

nal o

f Cel

l Sci

ence

ResultsVisualizing cortical actin cables by TIRFMTo visualize actin cables, we fused the endogenous copy of ABP140to green fluorescent protein (GFP) as previously reported (Yangand Pon, 2002). By employing TIRFM actin, structures labeled byAbp140–GFP could be visualized with improved sensitivity (Fig.1A). Due to the limited illumination depth of TIRFM (Axelrod etal., 1983), we had to ensure that actin cables were associated withthe cell cortex as previously reported (Amberg, 1998). We thereforeanalyzed Z-stacks of Abp140–GFP-labeled yeast cells using wide-field microscopy (Fig. 1B), and counted an average of 6.6±2.1(mean ± s.d., n51) cables for each mother cell or unbudded cellanalyzed. Cables in small and medium-sized buds were hardlyvisible and were therefore not included for quantitative analyses.Importantly, nearly all cables (n336) were found restricted to thecortical planes with only 1.2% extending into the interior of cells(Fig. 1B). Our results indicate that TIRFM is a reliable techniquefor monitoring actin cable dynamics in S. cerevisiae cells.

Actin cable dynamics are tightly regulated during cellpolarizationTo characterize cable dynamics in detail we followed individualcables moving through the field of view. Actin cables were rarelystatic and often either exhibited elongation (Fig. 2A; supplementary

material Movie 1) or translational motility (Fig. 2B; supplementarymaterial Movie 2). Due to the illumination limit of TIRFM and theoval shape of S. cerevisiae cells, only 50–70% of the surface areaof a cellular hemisphere could be visualized (field of view with 2–5 m diameter). However, most actin cables extended beyond thefield of view at one or both ends and it was often impossible todirectly distinguish elongation from translational motility.Following reliable fiduciary marks on cables was not feasiblebecause Abp140 has very low affinity to actin filaments andexchanges on cables within less than 200 milliseconds(supplementary material Fig. S1A) (Riedl et al., 2008). We therefore

1534 Journal of Cell Science 124 (9)

Fig. 1. Cortical actin cable network in S. cerevisiae cells. (A)Comparison ofwide-field and TIRF microscopy for visualization of Abp140–GFP labeledactin in wild-type yeast cells (strain JYY141). Cell periphery is marked bydotted lines. Time is given in seconds. (B)Example of a Z-stack used for cablecounts. Relative z-position given inm. Arrowheads indicate cables close tothe cell surface, asterisks indicate one cable found away from the cell surface.Scale bars: 2m.

Fig. 2. Actin cable dynamics during cell polarization. (A)Cable elongation(arrowheads show leading end; supplementary material Movie 1). Thekymograph was generated along the dotted lines. (B)Cable translationalmotility (arrowheads show leading end and asterisks the lagging end;supplementary material Movie 2). Scale bars: 2m. (C)Probability densitydistributions of cable movement rates in JYY141 wild-type unpolarized(unpol) and polarized (pol) cells (see Materials and Methods for details).(D)Stacked bar graphs of cable movement rates in wild-type cells. (E,F)Cablemovements in synchronized cells. Abp140–GFP-expressing cells (JYY207)were arrested for 3 hours and released into the cell cycle through washout ofmethionine. Cable movements were measured during arrest, 40–50 minutesafter release, and 80–90 minutes after release. Insets in E: typical cellmorphology in G1 arrest (a), 40-50 minutes after release (b) and 80–90minutes after release (c). (G)Cable number (N) per cell and per minute inwild-type (n20) and arrest strains (n>10); **P<0.01, ***P<0.0001 comparedwith unpolarized cells. Bar graphs show means + s.e.m.

Jour

nal o

f Cel

l Sci

ence

defined all extensions of leading cable ends as ‘cable movement’and attempted to separate kinetic subpopulations of cables bycareful quantification and statistics. To this end, we analyzed themeasured velocities by kernel density estimation and plotted theirprobability density functions. This approach allows a smoothedrepresentation of the raw data distribution and is considered superiorto histograms (see Materials and Methods for details). In addition,we recorded the number of cables moving through the field ofview as a measure for cable abundance and turnover in a cell.

In unbudded cells with no observable polarized cable arrays(referred to hereafter as unpolarized cells; supplementary materialFig. S1B) we observed rapid cable movement with velocities up to5 m/second with probability densities peaking around 1.2m/second and 2.2 m/second (Fig. 2C; supplementary materialFig. S2A). In small or medium-budded cells (bud diameter lessthan half of mother diameter, referred to hereafter as polarizedcells; supplementary material Fig. S1C), cable movement wasoverall slower with a single peak around 1 m/second (Fig. 2C;supplementary material Fig. S2A). For simplicity, cable movementmeasurements were separated into three velocity categories: slow,below 1 m/second; intermediate, between 1 and 2 m/second;and fast, above 2 m/second. These were plotted into stacked bargraphs and used to supplement the kernel density analyses. Inpolarized cells, a reduction of fast movements and increase in slowmovements was clearly observed (Fig. 2D). In addition, the numberof emerging cables was significantly reduced in polarized cells(10.7±3.5 cables/minute, mean ± s.d., n20) compared tounpolarized cells (24.5±5.8 cables/minute, n20; Fig. 2G). Thesedata suggested a decrease in cable dynamics upon cell polarization.

To verify this in a synchronous cell population undergoingpolarization, we introduced Abp140–GFP into a strain that couldbe arrested in the G1 phase of the cell cycle by cyclin depletion(Richardson et al., 1989; Wedlich-Soldner et al., 2004). Cabledynamics were then measured before and after synchronous releaseof cells from the G1 cell cycle arrest. Confirming our previousmeasurements, cable movement velocities and cable numbers werereduced as cells became polarized (Fig. 2E,F). This effect becamemost pronounced after 80 minutes when most cells had alreadyestablished polarity (Wedlich-Soldner et al., 2004) and formedsmall buds (Fig. 2E, insets).

Cortical Bni1 drives rapid assembly of unpolarized actincablesActin cables in S. cerevisiae were reported to depend on forminactivity and we confirmed by TIRFM that cables were absent incells lacking both formins (bnr1 bni1-11; supplementary materialFig. S3A) (Evangelista et al., 2002) or the essential formincofactor profilin (pfy1-4; supplementary material Fig. S3A)(Wolven et al., 2000). We then studied cable dynamics inindividual formin knockout mutants. In unpolarized bni1 cells,cable numbers were reduced by 90% (Fig. 3A). Interestingly, wealso observed a selective reduction of cable movement atintermediate velocities (Fig. 3B,C). On the other hand, inunpolarized bnr1 cells, cable numbers were only reduced by24% (Fig. 3A) and cable movement velocities remained largelyunchanged with a slight reduction of the slow population (Fig.3B,C). These results suggest that Bni1 generates most of thecables in unpolarized cells and regulates cable movement at

1535Actin cable dynamics in S. cerevisiae

Fig. 3. Cable dynamics inunpolarized cells of forminmutants. (A)Cable numbers informin mutants (n>20); **P<0.01,***P<0.0001 compared with controlcells. Bar graphs show means +s.e.m. (B,C)Cable movement incontrol (identical to ‘unpol’ in Fig.2C,D) with bni1 (JYY22) andbnr1 (JYY24) mutants. (D,E)Cablemovement in bnr1 cells expressingBni1-4PPT (JYY183, control) andbni1-1PPT (JYY189). (F)Dual-colorTIRFM showing an Abp140-(1-214)-mRFPRuby (Abp140RFP)-labeledactin cable (arrowheads) originatingfrom a cortical Bni1–GFP patch(asterisks). Images were acquiredfrom strain JYY144, and weredeconvolved to improvevisualization. Scale bar: 1m.(G,H)Cable movement rates inbnr1 cells treated with indicatedconcentrations of LatB.

Jour

nal o

f Cel

l Sci

ence

intermediate velocities between 1 and 2 m/second. Bnr1, bycontrast, plays a minor role at this stage and might be associatedwith slow movement below 1 m/second.

To mechanistically understand cable movement we wanted todirectly link cable movements at intermediate velocities to Bni1-driven actin polymerization. Both Bni1 and Bnr1 contain forminhomology 1 (FH1) and FH2 domains (Evangelista et al., 2002).FH2 binds to the barbed end of the actin filament, whereas FH1recruits profilin-bound actin via polyproline tracks (PPT) (Kovar,2006). It was previously reported that the number of PPTs intruncated constructs determined the rate of filament elongation byBni1 in vitro (Paul and Pollard, 2008). We attempted to reproducethis result in cells. Therefore, the genomic copy of BNI1 wasmodified in a bnr1 background to either contain four (wild type)or only one PPT. This modification did not affect cable numbers(Fig. 3A), in accordance with in vitro results where nucleationefficiency was unaffected (Paul and Pollard, 2008). However,compared to cells expressing Bni1-4PPT, we found a clear andselective shift of the velocity population associated with Bni1 incells expressing bni1-1PPT (Fig. 3D, peak from 1.4 m/second to1 m/second; Fig. 3E). This result confirms the dependence ofintermediate cable movement velocities on Bni1 and alsocorroborated the reduced polymerization rates with fewer PPTs invitro (Paul and Pollard, 2008).

Because actin cables were localized predominantly at the cellcortex (Fig. 1B), Bni1 had to also localize to the cell cortex toregulate cable dynamics. Indeed, previous data suggested thatformins were activated by membrane association (Dong et al.,2003). However, only cytosolic Bni1 dots had been observed inunpolarized cells so far (Buttery et al., 2007). No clear evidence ofcortical localization of Bni1 in these cells was reported. Takingadvantage of the increased sensitivity of our setup we nowmonitored Bni1–GFP localization in unpolarized cells by TIRFM.We found that Bni1–GFP transiently accumulated in cortical patchesthat were distinct from endocytic actin patches (supplementarymaterial Fig. S3B,C). Importantly, using dual-color TIRFM withBni1–GFP and Abp140–RFP we observed short actin cablesoriginating from cortical Bni1 patches (Fig. 3F). In summary, wepropose that cortical Bni1 contributes to cable dynamics inunpolarized cells by driving filament elongation at intermediatevelocities of 1–2 m/second.

Cortical Myo2 drives rapid cable motility in unpolarizedcellsRemarkably, none of the formin mutations resulted in a reductionof the cable population moving faster than 2 m/second (Fig. 3B–E), suggesting that these movements might occur independently ofactin polymerization. To verify this, we treated bnr1 cells withlatrunculin B (LatB), which specifically inhibits actinpolymerization by sequestering actin monomers (Ayscough et al.,1997). At increasing LatB concentrations, cables becamediminished until they disappeared completely in 16 M LatB(supplementary material Fig. S3D). In cells treated with 2–8 MLatB, actin cables became shorter and underwent rapid translationalmotility (supplementary material Movie 3). This movement mostlyoccurred at rates above 2 m/second (Fig. 3G,H). These resultsstrongly suggest that translational cable motility wasmechanistically distinct from formin-driven cable elongation andoccurred at higher speed. The initial double-peak distribution (Fig.2C) could have resulted from the combination of these mechanisms.

In S. cerevisiae the only previously reported process occurringat velocities above 2 m/second is intracellular transport by thetype V myosin Myo2 (Schott et al., 2002; Sheltzer and Rose,2009). Therefore, we monitored cable dynamics in unpolarizedcells of various myo2 mutants. The temperature-sensitive mutantmyo2-66 is defective in actin binding (Johnston et al., 1991),whereas the myo2-16 mutant (Schott et al., 1999) has a defect incargo binding. At restrictive temperature (36°C), cable numbers incells of either mutant were not affected (Fig. 4A). Strikingly,populations of cables moving above 2 m/second were stronglyreduced for both mutants (Fig. 4B–E). This became especiallyapparent in bni1 myo2-16 double mutant cells. In these cells, thecable number was reduced by 73% (Fig. 4A), similar to the bni1mutant (Fig. 3A). Notably, over 70% of the remaining cables nowmoved with velocities below 1 m/second (Fig. 4D,E), consistentwith the loss of both Bni1-mediated (1–2 m/second) and Myo2-mediated (above 2 m/second) movements. Moreover, cables inmyo2 mutant cells frequently became immobile for extendedperiods (Fig. 4H), compared to rapid continuous motility in thecontrol strain (Fig. 2B, supplementary material Movie 3).Disruption of any other S. cerevisiae myosin (Myo4, Myo1,Myo3+5) did not cause any reduction of cable movement inunpolarized cells (supplementary material Fig. S4A,B) indicatinga specific role of Myo2 in this process.

Both myo2-16 and myo2-66 alleles lead to cell lethality at therestrictive temperature (Johnston et al., 1991; Schott et al., 1999).To rule out secondary physiological defects as the cause of changesin cable dynamics, we wanted to directly link the motor activity ofMyo2 to cable motility. To this end, we measured cable dynamicsin Myo2 mutants containing different numbers of IQ motifs intheir neck domain. Previous study showed that stepwise reductionof IQ motifs from six (wild type) to zero might reduce myosin stepsize, causing progressive slow-down of Myo2-dependent transportof secretory vesicles (Schott et al., 2002). Consistently, afterreduction of IQ motifs, the population of cables moving at velocitiesabove 2 m/second was progressively reduced in our TIRFM assay(Fig. 4F,G).

Because actin cables localized predominantly close to the plasmamembrane (Fig. 1B) (Amberg, 1998), Myo2 had to be anchored atthe cell cortex to mediate cable movement. Using dual-colorTIRFM we showed that Myo2–GFP formed relatively static corticalMyo2 patches (Fig. 4I), which were distinct from actin patches(supplementary material Fig. S4C). These colocalized with actincables along the path of translational motility (Fig. 4J). Our resultsdemonstrate that Myo2 not only functions as a cargo motor usingactin cable as tracks, but also functions as the motor for translationalcable movement along the cell cortex.

Bnr1 and Bni1 govern actin cable dynamics in polarizedmother cellsWe identified Bni1 and Myo2 as molecules driving cable movementabove 1 m/second in unpolarized cells. What then mediated slowcable movement below 1 m/second? Our measurements showedonly a few cables in this range in unpolarized cells, and lack ofBnr1 specifically abolished these cables (Fig. 3B,C). By contrast,slower cable movements were much more prominent in polarizedmother cells, with half of the cables moving below 1 m/second(Fig. 2C–F). The activity profile of Bnr1 closely correlated withthe observed differences in cable movement [high activity inpolarized cells (Pruyne et al., 2004a), low activity in unpolarized

1536 Journal of Cell Science 124 (9)

Jour

nal o

f Cel

l Sci

ence

cells (Chesarone et al., 2009)]. We therefore suspected that Bnr1mediated cable movement below 1 m/second.

To test our hypothesis, we studied cable dynamics in polarizedmother cells of formin mutants. Deletion of Bni1 caused no changein cable numbers (Fig. 5A) but nearly completely abolished cablemovement above 1 m/second (Fig. 5B,C). Almost all cables werenow polarized towards the bud neck (Fig. 5D), where Bnr1 wasreportedly localized (Fig. 5E) (Buttery et al., 2007). Surprisingly,cable number and movement remained largely unaffected bydeletion of Bnr1 (Fig. 5A–C), but more than 40% of the cableswere now depolarized (Fig. 4D). These randomly oriented cablesmoved significantly faster than polarized cables (1.4±0.74m/second vs 0.94±0.6 m/second, P<0.001; Fig. 4F,G). Notably,rapid cable motility above 2 m/second was absent from allpolarized cells (Fig. 5B,C) indicating a negligible role of Myo2-driven cable movement. In agreement with this, Myo2–GFP was

mostly found at the bud tips of polarized cells (Fig. 5H) (Lillie andBrown, 1994), whereas cortical density of Myo2 patches wassignificantly reduced in polarized mother cell compared tounpolarized cells (Fig. 5I; P<0.0005).

In summary, our results suggest that cable dynamics in polarizedcells are predominantly mediated by the two formins Bni1 andBnr1. Bud-neck-confined Bnr1 was associated with elongation ofpolarized cables at rates below 1 m/second. Bni1 appeared todrive slow elongation of polarized cables (<1 m/second) as wellas elongation of unpolarized cables at intermediate speeds (1–2m/second). This model is consistent with the concentration ofBni1 at bud tips and necks of polarized cells (Fig. 5J) (Pruyne etal., 2004a) as well as transient localization of Bni1 on the mothercell cortex (Fig. S3B) (Buttery et al., 2007). Myo2-driven rapidcable motility (>2 m) plays almost no role in cable dynamics atthis stage.

1537Actin cable dynamics in S. cerevisiae

Fig. 4. Myo2 drives rapid cable motilityabove 2m/second. (A)Cable numbersin wild-type control cells and myo2mutants (n20); ***P<0.001 comparedwith control cells. Bar graphs show means+ s.e.m. (B,C)Cable movement rates inmyo2-66 (JYY32) and control (JYY141)cells. (D,E)Cable movement in myo2-16(JYY79), bni1 myo2-16 (JYY34) andcontrol (JYY75) cells. (F,G)Cablemovement in cells expressing Myo2-6IQ(JYY148), myo2-2IQ (JYY149) andmyo2-0IQ (JYY150). Cells in F and Gwere treated with 2M LatB to reducecontribution of formin activity. All datawere from unpolarized cells. (H)Actincable in a myo2-16 cell arrested for 9seconds. Arrowheads indicate cable ends.(I)TIRFM image of cortical Myo2–GFPpatches. Image stack was acquired instrain JYY8. Kymograph was generatedalong the dotted straight line with 6 pixelsline width. Cell periphery is marked by adotted line. (J)Motile actin cablescolocalize with multiple Myo2 patchesalong their paths. Images were acquiredfrom strain JYY133. Arrowheads andasterisks indicate Myo2 patches. Imageswere smoothed by Gaussian convolutionto assist visualization. All data wereacquired from unpolarized cells. Scalebar: 2m.

Jour

nal o

f Cel

l Sci

ence

Cable dynamics are important for correct cell polarizationPolarized transport into the bud requires stable arrays of actincables (Pruyne et al., 2004b). Here, we established quantitativelythat such arrays could be generated by oriented formation andimmobilization of actin cables. However, the advantage ofmaintaining rapidly moving and randomly oriented cables inunpolarized cells was less obvious because this high energy-consuming process occurs before any polarized growth. Our resultssuggested that cells switch between fast and slow cable dynamicsby activating Bnr1 at the bud neck. We therefore decided to observecells that are forced to exclusively rely on Bnr1-driven slow cabledynamics by either deleting the other formin or by removing theBnr1 inhibitor Bud14 (Chesarone et al., 2009).

One possible role of rapid and random cable dynamics is toenable efficient cell polarity establishment, a self-organizing process

that depends on rapid recycling of polarity regulators such as Rho-GTPase Cdc42 (Wedlich-Soldner and Li, 2004). Recycling ofCdc42 is mediated by a combination of actin- and guaninenucleotide dissociation inhibitor (GDI)-dependent pathways(Slaughter et al., 2009). We assayed cell polarization during budformation by monitoring the localization of GFP–Cdc42 insynchronized cell populations released from G1 cell cycle arrest(Wedlich-Soldner et al., 2004). To better discern defects in actin-mediated recycling, we deleted the S. cerevisiae GDI Rdi1, whichcan extract Cdc42 from membranes (Tiedje et al., 2008). We thentested the effect of Bni1 deletion on polarization. Although deletionof either Rdi1 or Bni1 led to no obvious defects, polarization wassignificantly delayed in the double mutant (Fig. 6A). In addition,5% of all cells exhibited a striking phenotype with split caps orbifurcated buds (Fig. 6B).

Bud14 was first identified as a factor regulating various aspectof cell morphogenesis during vegetative growth and mating.Deletion of Bud14 led to defects in cell growth and to elongatedcells that undergo pseudohyphal growth (Cullen and Sprague,2002). It was recently shown that Bud14 regulates Bnr1 bydisplacing it from the barbed ends of actin filaments (Chesarone etal., 2009). Without Bud14, Bnr1 was expected to remain attachedto newly generated actin cables. Here we show that, strikingly,Bnr1–GFP, which could not be detected on the cortex of controlunpolarized cells, formed polarized caps and dot-like corticalstructures in unbudded bud14 cells (Fig. 6C). This abnormallocalization was also reflected in reduced rates of cable movement,consistent with the predicted Bnr1-dependent slow cable dynamics(Fig. 6D,E). These results indicate that cells with constantlyactivated Bnr1 and, hence, abnormally slow actin dynamics are notable to become depolarized. These constitutively polarized cellscan only grow in a polarized manner throughout the cell cycle,resulting in abnormal cell morphology or pseudohyphal growth.

DiscussionIn this study, we provide insights into the mechanisms underlyingactin cable organization and dynamics via spatio-temporalpatterning of formins and myosin. In unpolarized G1 cells, actincables are randomly oriented and their polymerization is largelymediated by the cortical formin Bni1 (Fig. 6F). The second formin,Bnr1, plays only a minor role in unpolarized cells and is supposedlykept inactive by Bud14. Cortical type V myosin Myo2 contributesto actin dynamics by mediating rapid sliding of actin cables alongthe plasma membrane. In polarized cells cable dynamics aresignificantly slowed down. Cables become mostly oriented towardsthe bud neck, where they are polymerized by bud-neck-confinedBnr1. At the same time, Bni1 is recruited to the bud tip and budneck, where it also participates in the generation of polarized cablearray. Myo2 is depleted from the cell cortex and targeted to the budby polarized transport (Fig. 6F).

We show that cables undergo growth and motility with velocitiesup to 5 m/second. Noticeably, even the slower cable movementsin polarized cells were much faster than those reported previously(0.3–0.6 m/second) (Yang and Pon, 2002; Huckaba et al., 2006).The strain used in those previous studies also showed fastermovement in our hands (supplementary material Fig. S4A,B). Alikely reason for this apparent discrepancy is the lower sensitivityof wide-field microscopy compared to TIRFM (Fig. 1A). Mostcables seen by wide-field fluorescence were polarized cable bundles(Fig. 1B), whereas TIRFM imaging additionally revealed manyrapidly moving, unpolarized cables (Fig. 1A; supplementary

1538 Journal of Cell Science 124 (9)

Fig. 5. Formins Bnr1 and Bni1 govern cable dynamics in polarized cells.(A)Cable numbers in control, bni1 (JYY22) and bnr1 (JYY24) polarizedmother cells. (B,C)Cable movement rates in control (equal to ‘pol’ in Fig.2C,D), bni1 and bnr1 polarized mother cells. (D)Proportion of randomlyoriented cables that do not originate from the bud neck in polarized mothercells (control n20; bni1 n18; bnr1 n20); *P<0.05, ***P<0.0005compared with control cells. (E)Bud neck localization of Bnr1–GFP (JYY7).(F,G)Movement rates of randomly oriented vs polarized cables in polarizedcells of bnr1. (H)Concentration of Myo2–GFP at the bud tip of a medium-budded cell. Image was acquired in strain JYY8. (I)Density of Myo2–GFPpatches on the cell cortex in unpolarized (unpol) and polarized mother (pol)cells of control strain JYY8. Patch density was calculated using TIRFM bycounting the number of Myo2–GFP patches emerged per minute in a cell(unpol n21, pol n19). Patch number was then divided by the area of the fieldof view (inm2) to yield the density (***P<0.0005 compared withunpolarized cells). (J)Concentration of Bni1–GFP at both the bud tip and thebud neck of small and medium-budded cells (JYY6). Images in E, H and Jwere acquired using wide-field optics. Bar graphs show means + s.e.m. Scalebars: 2m.

Jour

nal o

f Cel

l Sci

ence

material Fig. S1B and Movies 2, 3). Also, the use of a highlysensitive EMCCD camera enabled us to achieve higher temporalresolution in our sequences. These allowed us to study cablesmoving faster than 1 m/second, which were probablyunderrepresented in previous studies.

It was previously shown that the two yeast formins, Bni1 andBnr1, differ in their interaction partners and localization patterns(Dong et al., 2003; Moseley and Goode, 2005; Buttery et al., 2007;Chesarone et al., 2009). We now present evidence suggesting thatthey also assemble kinetically separable cable populations in a cellcycle-dependent manner. Bni1 was the dominant player in cableassembly in unpolarized G1 cells (Fig. 3A–F), whereas Bnr1 onlybecame localized and/or activated during bud formation (Fig. 5A–E). Importantly, altered cable dynamics in several formin mutantssuggested that Bni1 might polymerize actin cables twice as fast asBnr1. This notion was supported by the velocity peaks at 1.2–1.5m/second in cells containing Bni1 as the only formin (Fig.3B,D,G), and the peaks appearing faithfully at 0.5–0.7 m/secondin bni1 or Bnr1 hyperactivation strains (Fig. 3B, Fig. 4D, Fig.6D). Another major difference was that Bnr1-generated cableswere uniformly polarized towards the bud neck – the place of theirorigin, whereas even in polarized cells the Bni1-nucleated cableswere frequently randomly oriented (Fig. 5D). This could beexplained by a higher affinity of Bnr1 to actin barbed ends,consistent with a tenfold higher affinity of Bnr1 in biochemicalexperiments (Moseley and Goode, 2005) as well as the presenceof Bni1 outside polarized sites (supplementary material Fig. S3B)

(Buttery et al., 2007). In summary, we identified the switch betweenthe two yeast formins as a major mechanism for regulation ofcable dynamics.

Our TIRFM-based cable dynamics measurements were sensitiveenough to detect the effect of specific manipulation of the Bni1-FH1 domain on actin polymerization in vivo. Previously, Paul andPollard demonstrated that by reducing the number of PPT fromfour to one, polymerization rate by Bni1 is reduced by about 50%(Paul and Pollard, 2008). We introduced these FH1 mutations intoendogenously expressed Bni1 and found a clear shift of Bni1-dependent cable movements from around 1.4 m/second to around1.0 m/second, with no major changes in Myo2-driven rapidmotility (Fig. 3D,E). Therefore the trend of the in vitromeasurements could be reproduced faithfully, indicating thatTIRFM-based observations of in vivo cable kinetics could beused to directly study the biochemical mechanisms of actinpolymerization.

Type V myosins are well known for their role in polarizedtransport of various cargos (Pruyne et al., 2004b; Trybus, 2008).We describe here a surprising novel role for Myo2 in moving actinfilaments along the cell cortex. In unpolarized cells, sliding ofcables by Myo2 allowed continuous movement and mixing ofcables along the periphery of the cell. To our knowledge, this isalso the first demonstration of a myosin moving actin filaments atthe cortex. In polarized cells, cables formed polarized arrays thatwere probably anchored to the bud neck and could not be movedby Myo2 (Fig. 5D). In this scenario, we expect myosin molecules

1539Actin cable dynamics in S. cerevisiae

Fig. 6. Actin cable dynamics is tightly coupled to cellpolarization. (A)Efficiency of cell polarization in cellsdefective in cable dynamics. Cell polarization was monitoredusing GFP–Cdc42. Cells are considered polarized if theycontain concentrated GFP fluorescence (polar caps) on the cellperiphery. Shown here are the fractions of polarized cells incontrol (RWS120), bni1 (JYY215), rdi1 (RWS150) andbni1 rdi1 (JYY216) strains at different times after releasefrom G1 arrest. Error bars show means ± s.d., n3 experimentwith 50 cells each. Cell periphery is marked by a dotted line.(B)Formation of a single polarization cap in an rdi1 cell andsplit caps/buds in bni1 rdi1 cells. (C)Localization of Bnr1–GFP in unbudded control (JYY7) and bud14 (JYY166) cells.(D,E)Cable movement rates in unbudded control (JYY141,equal to ‘unpol’ in Fig. 2C,D) and bud14 (JYY160) cells.(F)Schematic model for actin cable regulation by spatial-temporal patterning of Bni1, Bnr1 and Myo2 in unpolarized andpolarized S. cerevisiae cells. Scale bars: 2m.

Jour

nal o

f Cel

l Sci

ence

to be removed from the cortex (Fig. 5I) and move along the stablecables to the bud, where they can support polarized organelletransport (Pruyne et al., 2004b). This dual function of Myo2 as amotor that moves both cargos and tracks provides an efficientmechanism of adaptation to cell cycle-specific requirements.

Assembly and rapid reorganization of the actin cytoskeleton isclearly required for migration of amoeba and mammalian cells(Bretschneider et al., 2002; Lammermann et al., 2008). It is lessclear why S. cerevisiae, a unicellular fungus enclosed in a cellwall, expends so much effort on keeping actin cables highlydynamic in unpolarized cells. We have shown here that reductionof cable motility can have serious consequences for polarityestablishment. In cells (bni1) that rely on slower cable assemblydriven by Bnr1, polarization after G1 release was significantlydelayed (Fig. 6A) and these cells often could not properly focussecretion on a single polarization site (Fig. 6B). One possibleexplanation for this phenotype is that in bni1 cells all cablesgenerated during polarization became stably anchored to the septinring by Bnr1 (Pruyne et al., 2004a) and served as fixed vectors fordelivery of Cdc42-containing vesicles (Wedlich-Soldner et al.,2003). In cases where cables happened to lie on opposite sides ofthe septin ring, separate Cdc42 caps would be formed, giving riseto the observed split-bud phenotype. Deleting Bud14, a negativeregulator of Bnr1, caused a severe slow-down of cable dynamicsas well as untimely polarization of cells in G1 (Fig. 6C–E). Thisillustrates the problems associated with overly stable actin cables.Rapid reorganization of cables might be necessary to enable aprecise response during highly dynamic processes such as polarityestablishment (Wedlich-Soldner and Li, 2004; Brandman et al.,2005). Cells might not even be able to attain a non-polarized statewithout Bnr1-inactivation and, hence, rapid cable reorganization.

Materials and MethodsCell culture and yeast strainsUnless otherwise stated, all cells were haploid, grown at room temperature (24°C) andanalyzed from logarithmically growing cultures. Standard media and genomic manip-ulation procedures (Janke et al., 2004) were used. To generate bni1-1PPT, a fragmentof BNI1 (1927–5862) was cloned into plasmid pRS315. The FH1 domain (3687–4056)was then replaced by a fragment encoding 1PPT as described (Paul and Pollard, 2008),Plasmids with intact FH1 (4PPT) or 1PPT were linearized and integrated into the BNI1locus of the bnr1 strain. All strains used in this study are listed in supplementarymaterial Table S1.

Plasmid construction for actin cable markersTo label actin cables with RFP we used the construct pRS315-Abp140(1-214)-mRFPruby, based on the pRS315 (CEN, LEU2) plasmid. It contains the Abp140promoter followed by the coding region for the N-terminal 214 amino acids of Abp140,which was fused to the mRFPruby gene (Fischer et al., 2006). The mRFPruby genewas amplified with primers 5�-GGATCCGGGTGCTGGTGCTGGTGCTGGTGC-TATGGGCAAGCTTACCATGGCCAG-3� and 5�-GCGGCCGCTTAAGCGCCTGT-GCTATGTCTGCC-3�. Expression of Abp140 (1-214)-mRFPruby from this plasmidlabels actin more strongly than endogenous Abp140, allowing the detection of cablesin two color experiments.

MicroscopyAll images were acquired on a custom TIRFM setup (Till Photonics, Gräfelfing,Germany) based on an automated iMic-stand and an Olympus 1.45 NA 100�objective. Lasers at 488 nm (Coherent Sapphire 75 mW) and 561 nm (Cobolt Jive75 mW) were coupled through an AOTF and a galvanometer-driven scan head to adjustTIRF angles. Images were collected with an Andor iXON DU-897 EMCCD cameraat maximum gain after a 2� magnifying lens (Andor, Belfast, N. Ireland). Acquisitionwas controlled by the Live Acquisition software package. Wide-field images wereacquired using the same setup using 0° incidence angle for the laser. Z-sectioning wasperformed using the built-in nanodrive system in the iMic setup. The epifluorescencemicroscope used for counting cables (Fig. 1B) was a Zeiss Axio Imager A1 with a Zeiss100� 1.45 NA objective coupled to an ASI MS-2000 XYZ stage. Images werecollected with an Andor iXON DU-897 EMCCD camera after a 2� magnifying lens(Zeiss) and Metamorph software.

Drug treatment and G1 cell cycle arrestCells subjected to LatB treatment were grown in drug-containing media at roomtemperature for 30–45 minutes prior to imaging. For G1 arrest assay, cln-arrest cells(Richardson et al., 1989) were grown to OD 0.5–0.8 and arrested in synthetic dropoutmedium containing 3 mM methionine for 2–3 hours. Cells were then released bywashing out the methionine. Cable dynamics were imaged before and after release inat least 10 cells per time point. Cell polarization was monitored by determining thepercentage of cells exhibiting concentration polar cap of GFP–Cdc42 fluorescence onthe plasma membrane.

Measurement of cable movementImages were acquired in TIRF mode at 5 Hz. Cable ends were traced over at least twoframes. Distance of cable movement was divided by elapsed time to obtain velocities.

Kernel density estimationThis technique is a non-parametric way of estimating the probability density functionof a random variable and allows smoothing of a finite data set. It avoids the biasinherent in histograms when selecting bin size and endpoints. Kernel estimatorssmooth out the contribution of each observed data point over a local neighborhoodof that data point. Here raw measurements of cable velocities were processed andplotted using the Gaussian kernel density function in MATLAB (The MathWorks),given as:

where p(x) is the probability density of events at a given velocity, n is the samplesize and h is the bandwidth of the kernel. The bandwidth was estimated for eachdataset as:

where is the estimated standard deviation of the dataset and Q.75 and Q.25 are its75% and 25% quartiles, respectively.

Image processingUnless specified otherwise, images are raw or inverted for visualization. Forsmoothening, images were convolved with a Gaussian kernel (1 3 1; 3 7 3; 1 3 1).Iterative deconvolution was performed in Huygens (Scientific Volume Imaging,Hilversum, The Netherlands). Point spread functions were calculated separately for488 nm and 561 nm lasers, using 40 nm yellow–green fluorescent microbeads(Invitrogen).

StatisticsAll averages are given as geometric mean ± s.e.m. or s.d. P-values were calculatedusing the t-test (Mann-Whitney) in Prism (GraphPad).

We thank Anthony Bretscher (Cornell University, Ithaca, NY), LizaPon (Columbia University Medical Center, New York, NY) and RongLi (Stowers Institute for Medical Research, Kansas City, MO) forplasmids and strains, S. Wedlich, S. Oliferenko, T. Soldati and membersof the Wedlich-Soldner group for comments and discussion on themanuscript, and G. Beck for technical support. This work was supportedby the Max-Planck Society.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/124/9/1533/DC1

ReferencesAmberg, D. C. (1998). Three-dimensional imaging of the yeast actin cytoskeleton through

the budding cell cycle. Mol. Biol. Cell 9, 3259-3262.Axelrod, D., Thompson, N. L. and Burghardt, T. P. (1983). Total internal inflection

fluorescent microscopy. J. Microsc. 129, 19-28.Ayscough, K. R., Stryker, J., Pokala, N., Sanders, M., Crews, P. and Drubin, D. G.

(1997). High rates of actin filament turnover in budding yeast and roles for actin inestablishment and maintenance of cell polarity revealed using the actin inhibitorlatrunculin-A. J. Cell Biol. 137, 399-416.

Brandman, O., Ferrell, J. E., Jr, Li, R. and Meyer, T. (2005). Interlinked fast and slowpositive feedback loops drive reliable cell decisions. Science 310, 496-498.

Bretschneider, T., Jonkman, J., Kohler, J., Medalia, O., Barisic, K., Weber, I., Stelzer,E. H., Baumeister, W. and Gerisch, G. (2002). Dynamic organization of the actinsystem in the motile cells of Dictyostelium. J Muscle Res. Cell Motil. 23, 639-649.

Buttery, S. M., Yoshida, S. and Pellman, D. (2007). Yeast formins Bni1 and Bnr1 utilizedifferent modes of cortical interaction during the assembly of actin cables. Mol. Biol.Cell 18, 1826-1838.

Chalkia, D., Nikolaidis, N., Makalowski, W., Klein, J. and Nei, M. (2008). Origins andevolution of the formin multigene family that is involved in the formation of actinfilaments. Mol. Biol. Evol. 25, 2717-2733.

h = 1.06 min σ ,Q.75− Q.25

1.34

⎛⎝⎜

⎞⎠⎟

n−

1

5 , (2)

p(x) =1

nh 2πe

−x − xi

2h

⎛

⎝⎜

⎞

⎠⎟

2

i =1

n

∑ , (1)

1540 Journal of Cell Science 124 (9)

Jour

nal o

f Cel

l Sci

ence

Chesarone, M., Gould, C. J., Moseley, J. B. and Goode, B. L. (2009). Displacement offormins from growing barbed ends by bud14 is critical for actin cable architecture andfunction. Dev. Cell 16, 292-302.

Clayton, J. E., Sammons, M. R., Stark, B. C., Hodges, A. R. and Lord, M. (2010).Differential regulation of unconventional fission yeast myosins via the actin track. Curr.Biol. 20, 1423-1431.

Cullen, P. J. and Sprague, G. F., Jr (2002). The Glc7p-interacting protein Bud14pattenuates polarized growth, pheromone response, and filamentous growth inSaccharomyces cerevisiae. Eukaryot. Cell 1, 884-894.

Diez, S., Gerisch, G., Anderson, K., Muller-Taubenberger, A. and Bretschneider, T.(2005). Subsecond reorganization of the actin network in cell motility and chemotaxis.Proc. Natl. Acad. Sci. USA 102, 7601-7606.

Dong, Y., Pruyne, D. and Bretscher, A. (2003). Formin-dependent actin assembly isregulated by distinct modes of Rho signaling in yeast. J. Cell Biol. 161, 1081-1092.

Evangelista, M., Pruyne, D., Amberg, D. C., Boone, C. and Bretscher, A. (2002).Formins direct Arp2/3-independent actin filament assembly to polarize cell growth inyeast. Nat. Cell Biol. 4, 32-41.

Fischer, M., Haase, I., Wiesner, S. and Muller-Taubenberger, A. (2006). Visualizingcytoskeleton dynamics in mammalian cells using a humanized variant of monomericred fluorescent protein. FEBS Lett. 580, 2495-2502.

Geli, M. I. and Riezman, H. (1996). Role of type I myosins in receptor-mediatedendocytosis in yeast. Science 272, 533-535.

Higashida, C., Miyoshi, T., Fujita, A., Oceguera-Yanez, F., Monypenny, J., Andou, Y.,Narumiya, S. and Watanabe, N. (2004). Actin polymerization-driven molecularmovement of mDia1 in living cells. Science 303, 2007-2010.

Huckaba, T. M., Lipkin, T. and Pon, L. A. (2006). Roles of type II myosin and atropomyosin isoform in retrograde actin flow in budding yeast. J. Cell Biol. 175, 957-969.

Huh, W. K., Falvo, J. V., Gerke, L. C., Carroll, A. S., Howson, R. W., Weissman, J. S.and O’Shea, E. K. (2003). Global analysis of protein localization in budding yeast.Nature 425, 686-691.

Janke, C., Magiera, M. M., Rathfelder, N., Taxis, C., Reber, S., Maekawa, H., Moreno-Borchart, A., Doenges, G., Schwob, E., Schiebel, E. et al. (2004). A versatile toolboxfor PCR-based tagging of yeast genes: new fluorescent proteins, more markers andpromoter substitution cassettes. Yeast 21, 947-962.

Jansen, R. P., Dowzer, C., Michaelis, C., Galova, M. and Nasmyth, K. (1996). Mothercell-specific HO expression in budding yeast depends on the unconventional myosinmyo4p and other cytoplasmic proteins. Cell 84, 687-697.

Johnston, G. C., Prendergast, J. A. and Singer, R. A. (1991). The Saccharomycescerevisiae MYO2 gene encodes an essential myosin for vectorial transport of vesicles.J. Cell Biol. 113, 539-551.

Karpova, T. S., McNally, J. G., Moltz, S. L. and Cooper, J. A. (1998). Assembly andfunction of the actin cytoskeleton of yeast: relationships between cables and patches. J.Cell Biol. 142, 1501-1517.

Kovar, D. R. (2006). Molecular details of formin-mediated actin assembly. Curr. Opin.Cell Biol. 18, 11-17.

Kovar, D. R. and Pollard, T. D. (2004). Progressing actin: formin as a processiveelongation machine. Nat. Cell Biol. 6, 1158-1159.

Lammermann, T., Bader, B. L., Monkley, S. J., Worbs, T., Wedlich-Soldner, R.,Hirsch, K., Keller, M., Forster, R., Critchley, D. R., Fassler, R. et al. (2008). Rapidleukocyte migration by integrin-independent flowing and squeezing. Nature 453, 51-55.

Lillie, S. H. and Brown, S. S. (1994). Immunofluorescence localization of theunconventional myosin, Myo2p, and the putative kinesin-related protein, Smy1p, to thesame regions of polarized growth in Saccharomyces cerevisiae. J. Cell Biol. 125, 825-842.

McConnell, R. E. and Tyska, M. J. (2007). Myosin-1a powers the sliding of apicalmembrane along microvillar actin bundles. J. Cell Biol. 177, 671-681.

Moseley, J. B. and Goode, B. L. (2005). Differential activities and regulation ofSaccharomyces cerevisiae formin proteins Bni1 and Bnr1 by Bud6. J. Biol. Chem. 280,28023-28033.

Moseley, J. B. and Goode, B. L. (2006). The yeast actin cytoskeleton: from cellularfunction to biochemical mechanism. Microbiol. Mol. Biol. Rev. 70, 605-645.

Paul, A. S. and Pollard, T. D. (2008). The role of the FH1 domain and profilin in formin-mediated actin-filament elongation and nucleation. Curr. Biol. 18, 9-19.

Paul, A. S. and Pollard, T. D. (2009). Review of the mechanism of processive actinfilament elongation by formins. Cell Motil. Cytoskeleton 66, 606-617.

Pruyne, D. W., Schott, D. H. and Bretscher, A. (1998). Tropomyosin-containing actincables direct the Myo2p-dependent polarized delivery of secretory vesicles in buddingyeast. J. Cell Biol. 143, 1931-1945.

Pruyne, D., Gao, L., Bi, E. and Bretscher, A. (2004a). Stable and dynamic axes ofpolarity use distinct formin isoforms in budding yeast. Mol. Biol. Cell 15, 4971-4989.

Pruyne, D., Legesse-Miller, A., Gao, L., Dong, Y. and Bretscher, A. (2004b). Mechanismsof polarized growth and organelle segregation in yeast. Annu. Rev. Cell Dev. Biol. 20,559-591.

Richardson, H. E., Wittenberg, C., Cross, F. and Reed, S. I. (1989). An essential G1function for cyclin-like proteins in yeast. Cell 59, 1127-1133.

Riedl, J., Crevenna, A. H., Kessenbrock, K., Yu, J. H., Neukirchen, D., Bista, M.,Bradke, F., Jenne, D., Holak, T. A., Werb, Z. et al. (2008). Lifeact: a versatile markerto visualize F-actin. Nat. Methods 5, 605-607.

Sagot, I., Klee, S. K. and Pellman, D. (2002). Yeast formins regulate cell polarity bycontrolling the assembly of actin cables. Nat. Cell Biol. 4, 42-50.

Schott, D., Ho, J., Pruyne, D. and Bretscher, A. (1999). The COOH-terminal domain ofMyo2p, a yeast myosin V, has a direct role in secretory vesicle targeting. J. Cell Biol.147, 791-808.

Schott, D. H., Collins, R. N. and Bretscher, A. (2002). Secretory vesicle transportvelocity in living cells depends on the myosin-V lever arm length. J. Cell Biol. 156, 35-39.

Sheltzer, J. M. and Rose, M. D. (2009). The class V myosin Myo2p is required for Fus2ptransport and actin polarization during the yeast mating response. Mol. Biol. Cell 20,2909-2919.

Skau, C. T. and Kovar, D. R. (2010). Fimbrin and tropomyosin competition regulatesendocytosis and cytokinesis kinetics in fission yeast. Curr. Biol. 20, 1415-1422.

Slaughter, B. D., Das, A., Schwartz, J. W., Rubinstein, B. and Li, R. (2009). Dualmodes of cdc42 recycling fine-tune polarized morphogenesis. Dev. Cell 17, 823-835.

Sousa, A. D. and Cheney, R. E. (2005). Myosin-X: a molecular motor at the cell’sfingertips. Trends Cell Biol. 15, 533-539.

Staiger, C. J., Sheahan, M. B., Khurana, P., Wang, X., McCurdy, D. W. and Blanchoin,L. (2009). Actin filament dynamics are dominated by rapid growth and severing activityin the Arabidopsis cortical array. J. Cell Biol. 184, 269-280.

Tiedje, C., Sakwa, I., Just, U. and Hofken, T. (2008). The Rho GDI Rdi1 regulates RhoGTPases by distinct mechanisms. Mol. Biol. Cell 19, 2885-2896.

Trybus, K. M. (2008). Myosin V from head to tail. Cell. Mol. Life Sci. 65, 1378-1389.Wang, Y. L. (2001). The mechanism of cytokinesis: reconsideration and reconciliation.

Cell Struct. Funct. 26, 633-638.Wedlich-Soldner, R. and Li, R. (2004). Closing the loops: new insights into the role and

regulation of actin during cell polarization. Exp. Cell Res. 301, 8-15.Wedlich-Soldner, R., Altschuler, S., Wu, L. and Li, R. (2003). Spontaneous cell

polarization through actomyosin-based delivery of the Cdc42 GTPase. Science 299,1231-1235.

Wedlich-Soldner, R., Wai, S. C., Schmidt, T. and Li, R. (2004). Robust cell polarity isa dynamic state established by coupling transport and GTPase signaling. J. Cell Biol.166, 889-900.

Wolven, A. K., Belmont, L. D., Mahoney, N. M., Almo, S. C. and Drubin, D. G. (2000).In vivo importance of actin nucleotide exchange catalyzed by profilin. J. Cell Biol. 150,895-904.

Wu, J. Q., Sirotkin, V., Kovar, D. R., Lord, M., Beltzner, C. C., Kuhn, J. R. andPollard, T. D. (2006). Assembly of the cytokinetic contractile ring from a broad bandof nodes in fission yeast. J. Cell Biol. 174, 391-402.

Yanagida, T. (2007). Muscle contraction mechanism based on actin filament rotation. Adv.Exp. Med. Biol. 592, 359-367.

Yang, H. C. and Pon, L. A. (2002). Actin cable dynamics in budding yeast. Proc. Natl.Acad. Sci. USA 99, 751-756.

Zot, H. G. and Pollard, T. D. (1993). Motility of myosin I on planar lipid surfaces.Methods Cell Biol. 39, 51-63.

1541Actin cable dynamics in S. cerevisiae

Jour

nal o

f Cel

l Sci

ence