the yeast actin cytoskeleton: from cellular …teins and their biochemical mechanisms of action. for...

MICROBIOLOGY AND MOLECULAR BIOLOGY REVIEWS, Sept. 2006, p. 605–645 Vol. 70, No. 31092-2172/06/$08.00�0 doi:10.1128/MMBR.00013-06Copyright © 2006, American Society for Microbiology. All Rights Reserved.

The Yeast Actin Cytoskeleton: from Cellular Functionto Biochemical MechanismJames B. Moseley and Bruce L. Goode*

Department of Biology and The Rosenstiel Basic Medical Sciences Research Center,Brandeis University, Waltham, Massachusetts 02454

INTRODUCTION .......................................................................................................................................................606ACTIN STRUCTURES AND FUNCTIONS IN YEAST CELLS..........................................................................606

First Contact............................................................................................................................................................606Actin Patches and Endocytosis .............................................................................................................................607

Early observations ..............................................................................................................................................607Actin patch dynamics .........................................................................................................................................607Actin patch ultrastructure .................................................................................................................................609Unified model for actin patch development and function ............................................................................609

(i) Early recruitment: nonmotile phase .......................................................................................................609(ii) Intermediate stage in patch development: slow motility ....................................................................611(iii) Scission and rapid transport of patches .............................................................................................611

Actin Cables: Dynamic Tracks for Polarized Growth .......................................................................................611Introduction .........................................................................................................................................................611Mechanism of actin cable formation ...............................................................................................................612The bud tip polarity cap ....................................................................................................................................613Actin cable architecture .....................................................................................................................................613Gone in 60 seconds: actin cables are composed of short filaments ............................................................614Actin cable dynamics and turnover..................................................................................................................615

Yeast Cytokinesis: Two Complementary Mechanisms for Cell Division ........................................................616Introduction .........................................................................................................................................................616Targeting secretion to the bud neck at cytokinesis........................................................................................617Formation and closure of the actomyosin ring ..............................................................................................617Role of yeast myosin light chains in cytokinesis ............................................................................................618Coordinating septum formation with actin ring contraction .......................................................................618Septins ..................................................................................................................................................................618

MECHANISMS OF ACTIN FILAMENT ASSEMBLY, ORGANIZATION, AND TURNOVER .....................619Introduction .............................................................................................................................................................619

Biochemical properties of actin ........................................................................................................................619Assembly of Actin at Cortical Patches.................................................................................................................619

The Arp2/3 complex............................................................................................................................................619NPFs .....................................................................................................................................................................621

(i) Las17/WASp ...............................................................................................................................................623(ii) Pan1 ...........................................................................................................................................................624(iii) Myo3 and Myo5.......................................................................................................................................624(iv) Abp1 ..........................................................................................................................................................624

Coronin: spatial inhibitor of Arp2/3 complex activity...............................................................................................625Open questions about the Arp2/3 complex .....................................................................................................625

Assembly of Actin Cables ......................................................................................................................................625Formins Bni1 and Bnr1 .....................................................................................................................................625Profilin-FH1 interactions: a throttle for filament elongation.......................................................................627Bud6: stimulation of Bni1-mediated actin assembly .....................................................................................628Rho GTPases .......................................................................................................................................................628Other Bni1 and Bnr1 ligands ...........................................................................................................................629

Organization and Stabilization of Actin Filaments ...........................................................................................629Bundling proteins ...............................................................................................................................................629

(i) Sac6 .............................................................................................................................................................629(ii) Scp1............................................................................................................................................................630(iii) Iqg1/Cyk1 .................................................................................................................................................630(iv) Crn1...........................................................................................................................................................631(v) Abp140........................................................................................................................................................631

* Corresponding author. Mailing address: Rosenstiel Center, Bran-deis University, 415 South Street, Waltham, MA 02454. Phone: (781)736-2464. Fax: (781) 736-2405. E-mail: [email protected].

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(vi) Tef1 and Tef2 ...........................................................................................................................................631Nonbundling Actin Stabilizers ..............................................................................................................................632

(i) Tropomyosins Tpm1 and Tpm2 ..............................................................................................................632(ii) Capping protein........................................................................................................................................632

Rapid Turnover of Actin Structures ....................................................................................................................633Importance of actin turnover in vivo ...............................................................................................................633Cofilin/ADF ..........................................................................................................................................................633Aip1.......................................................................................................................................................................634Srv2/CAP ..............................................................................................................................................................634Profilin..................................................................................................................................................................635Twinfilin ...............................................................................................................................................................636

CLOSING REMARKS AND FUTURE DIRECTIONS .........................................................................................637ACKNOWLEDGMENTS ...........................................................................................................................................637REFERENCES ............................................................................................................................................................637

INTRODUCTION

All cells undergo rapid remodeling of their actin networks toregulate such critical processes as endocytosis, cytokinesis, cellpolarity, and cell morphogenesis. These events are driven bythe coordinated activities of a set of 20 to 30 highly conservedactin-associated proteins, in addition to many cell-specific ac-tin-associated proteins and numerous upstream signaling mol-ecules. The combined activities of these factors control withexquisite precision the spatial and temporal assembly of actinstructures and ensure dynamic turnover of actin structuressuch that cells can rapidly alter their cytoskeletons in responseto internal and external cues. One of the most exciting princi-ples to emerge from the last decade of research on actin is thatthe assembly of architecturally diverse actin structures is gov-erned by highly conserved machinery and mechanisms. Withthis realization, it has become apparent that pioneering effortsin budding yeast have contributed substantially to defining theuniversal mechanisms regulating actin dynamics in eukaryotes.

In this review, we first describe the filamentous actin (F-actin) structures found in Saccharomyces cerevisiae (patches,cables, and rings) and their physiological functions, and thenwe discuss in detail the specific roles of actin-associated pro-teins and their biochemical mechanisms of action. For moredetailed information on actin-based cellular processes, we re-fer readers to a number of excellent recent reviews that coverthese topics in greater depth: endocytosis (93), mitochondrialinheritance (42), vacuolar inheritance (45), establishment ofcell polarity (302, 305, 334), and cytokinesis (22, 33). In somesections we present data and concepts in a historical order,where tracing the progression is instructive. We also note thatthe majority of the cellular actin structures and their compo-nents are conserved between budding yeast and fission yeast,and in many cases insights from fission yeast preceded thosefrom budding yeast. Due to space constraints we have focusedthe review on the budding yeast actin cytoskeleton, but wehighlight experiments from fission yeast to reinforce specificpoints.

ACTIN STRUCTURES AND FUNCTIONSIN YEAST CELLS

First Contact

The first major insights into organization of the yeast actincytoskeleton came from two landmark papers in 1984 by

Adams, Kilmartin, and Pringle (2, 183). In the preceding years,actin had been purified from yeast cells (393), and it had beenestablished that actin was expressed from a single essentialgene in S. cerevisiae, ACT1 (349), but the distribution of actinin yeast cells remained a mystery. Using fluorochrome-labeleddrugs (2) and actin antibodies (183), Adams and coworkersvisualized filamentous actin in chemically fixed cells and de-scribed an “unusual distribution of actin” consisting of twodistinct structures (Fig. 1): cortical spots (“patches”) and longfibers (“cables”). Their elegant description of actin organiza-tion (see details below) set the stage for the recent advances incell biological analyses on yeast actin described in the followingsections: “Numerous small patches or spots were seen distrib-uted over the cell surface; the cortical location of these spots

FIG. 1. Cell cycle-regulated organization of the S. cerevisiae actincytoskeleton. Yeast cells at different stages in the cell cycle containthree visible F-actin structures: cortical actin patches, polarized actincables, and a cytokinetic actin ring. While patches and cables arevisible throughout the cell cycle, the ring is visible shortly before andduring cytokinesis. Shown here are actively growing cells from anasynchronous culture that were chemically fixed and stained with rho-damine phalloidin to visualize filamentous actin structures. (Modifiedfrom reference 12 with permission of the publisher.)

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was clear when the microscope was focused up and down. Inaddition, fibers were seen coursing through the cells, generallyin a direction roughly parallel to the long axes of the cells. Inunbudded cells, there was often a concentration of fluores-cence near one pole, and in cells with very small buds there wasgenerally a pronounced concentration of fluorescence in thebud. In both cases, the concentration of fluorescence was usu-ally resolvable as a cluster of individual spots. In cells withmedium-sized buds, there was typically a higher concentrationof fluorescent spots in the bud than in the mother cell. Cellswith very large buds often showed a concentration of fluores-cence in the neck region, sometimes apparently as clusters ofspots and sometimes apparently as a band.” (Reproduced withpermission from reference 2.)

In addition to patches and cables, Adams and coworkersnoted the presence of a third actin structure located at theneck of some large-budded cells, which they referred to(above) as a “band.” Twenty years later, it is clear that this wasthe first sighting of the budding yeast actomyosin contractilering. However, the concept of a budding yeast actomyosin ringlay relatively dormant over the next 14 years, in part due to thefact that actin rings are present only transiently during the cellcycle and therefore observed in only a limited number of cellsin an asynchronously growing culture and in part because theycan be more difficult to image than patches and cables. Inaddition, many in the field believed that the narrowness of thebud neck, approximately 1 �m during budding yeast cytokine-sis (327), precluded the requirement for an actomyosin con-tractile ring. Then, in 1998, the presence of a functional acto-myosin ring was confirmed by two independent studies (34,217) which revealed that the events of cytokinesis in yeast andanimals were more similar than previously thought.

Studies in 1986 revealed that the three visible F-actin struc-tures in budding yeast—patches, cables, and rings—are alsopresent in the fission yeast Schizosaccharomyces pombe (231).A large body of literature since then has shown that the mech-anisms regulating patch and cable assembly in fission and bud-ding yeast are similar but that the rules governing assembly ofthe actomyosin ring differ significantly (see below). In addition,smaller and/or less visible actin structures may exist in bothfission and budding yeast, as suggested by the requirement foractin dynamics in the processes of vacuolar fusion (92), endo-plasmic reticulum cortical dynamics (301), and chromatin re-modeling (38). Visualizing these actin structures in vivo rep-resents a key challenge for the future.

Actin Patches and Endocytosis

Early observations. Initial observations showing that patchesare concentrated at sites of polarized growth suggested thatpatch function might be linked to secretion. However, patchesare decorated with numerous actin-associated proteins, and asmutations in these patch components were characterized, itbecame apparent that their primary defects were endocyticrather than secretory. Multiple patch components (e.g., actin,Arc35, Rvs167, Sac6, Sla2, and Vrp1) were identified in ran-dom screens for mutants defective in endocytosis, end mutants(64, 86, 93, 202, 260, 309, 311). In addition, an endocyticfunction for actin was suggested by defects in fluid uptake(marked by Lucifer Yellow) upon treatment of cells with

latrunculin-A (Lat-A), an actin monomer-sequestering agent(19). Over the following years, genetic studies continued tosteadily draw correlations between the functions of patch com-ponents and endocytosis, but this linkage was not solidified andfully accepted until recent light and electron microscopy (EM)studies addressed the dynamic movement and ultrastructure ofcortical actin patches (see below).

Although it is now apparent that actin patches mediate en-docytosis, they may also be coupled to exocytosis. No patchcomponents were isolated in the original sec mutant screen(271), but mutants of a number of patch components show anaccumulation of post-Golgi vesicles (139, 213, 257), consistentwith models that suggest temporal and spatial links betweenendocytosis and exocytosis.

Actin patch dynamics. An important and unanticipatedproperty of actin patches was discovered with the advent ofgreen fluorescent protein (GFP) fusions to monitor real-timebehavior of proteins in cells. By tagging patch components(Sac6, Cap1/Cap2, Abp1, and actin) with GFP, two initial stud-ies demonstrated that patches are highly motile (80, 387).These and other early studies showed that the lifetime of actinpatches is approximately 10 to 20 seconds and that actinpatches first assemble at sites of polarized cell growth and thenmove slowly and nondirectionally along the cell cortex. Inaddition, more-rapid movements were observed. Patch motilityrates ranged from 0.1 to 0.5 �m/s (52, 354, 387).

These observations raised an important mechanistic ques-tion: what provides the force driving patch movement? A de-cade ago, when patch motility was first observed, most forms ofactin-based motility were thought to be myosin dependent. So,it came as a surprise when it was reported that the rate of patchmotility was unaffected by mutations in any of the five yeastmyosin genes: MYO3 and MYO5 (type I), MYO1 (type II), orMYO2 and MYO4 (type V) (354, 387). Instead, studies usingLat-A suggested that the actin filaments in patches undergorapid turnover (19) and that Lat-A inhibits patch movement(52, 289), suggesting that actin polymerization may provide theforce required for patch motility. As such, parallels were drawnbetween yeast actin patch motility and the actin polymeriza-tion-based motility of the intracellular pathogen Listeria mono-cytogenes, which hijacks the host actin nucleation machinery(the Arp2/3 complex) to assemble a highly branched actin“comet tail” (287, 407), propelling the bacterium at approxi-mately 0.4 �m/s (70, 368). While it has been appealing inmodels to depict actin patches like miniature Listeria actincomet tails trailing endocytic vesicles, patches have at least twokey properties that distinguish them from Listeria. First, cofilinactivity, and therefore rapid actin turnover, is required forListeria motility (287) but not rapid patch movement (204,287). Second, Listeria motility is autonomous in living cells andcell extracts, requiring only the actin tail that it forms forpropulsion. However, the rapid phase of patch motility (seebelow) relies on an additional separate network of filamentousactin structures, actin cables (159).

From a number of recent studies, it has become apparentthat actin patches mature in stages corresponding to differentstages of endocytosis and characterized by different types ofmovement. One of the first indications of this behavior camefrom a study examining the dual localization and fluorescenceresonance energy transfer between CFP-Abp1 (an activator of

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the Arp2/3 complex) and YFP-Sla1 (an early endocytic pro-tein) (391). This showed that at any given time, a subset ofpatches colabeled with Abp1 and Sla1, while others containedexclusively Sla1 or Abp1. Intriguingly, the patches labeled ex-clusively with Abp1 were highly motile, whereas those labeledexclusively with Sla1 exhibited little movement. The authorshypothesized that “Sla1-only” patches might contain endocyticmachinery and the arrival of Abp1 (and possibly other factors)facilitated actin polymerization-based rapid movement ofpatches. This study emphasized the need to consider temporalchanges in the development and lifetime of an actin patch.

Subsequent studies by Kaksonen et al. (173, 174) resolvedmany of the key events in patch development by elegantlycorrelating temporal changes in patch motility with the arrivalof specific components. Using pairs of integrated functionalfluorescent tags on different patch components (Las17, Sla1,Sla2, Pan1, Abp1, and the Arp2/3 complex) and computeralgorithms to track patch movements, they defined three stagesin the lifetime of a patch. First, nonmotile patches, whichcontain Las17, Sla1, and Pan1, but not actin, form at the cellcortex. Next, polymerized actin appears, which coincides withthe onset of slow patch movements at the cortex (0.05 to 0.1�m/s). Patches transition to a phase of rapid inward movementfrom the cell cortex, likely as vesicles coated with actin fila-ments. Addition of Lat-A abolishes both the slow and fastpatch movements (stages 2 and 3), suggesting that this motilityis actin polymerization based (289, 407). The roles of specificpatch components are discussed below in a model for patchdevelopment.

Shortly after the initial Kaksonen study (173), Huckaba et al.(159) reported three key findings clarifying patch function andmechanism. First, they showed that endocytic vesicles (markedby the lipophilic marker FM4-64) colocalize with motilepatches (marked by Abp1-GFP) during slow and fast phases ofmovement (Fig. 2A). This provided direct evidence that actinpatches are sites of endocytic uptake. Second, they showed thatrapid actin patch movement is mediated by polarized actincables, as internalized actin patches colocalize with cables (Fig.2B) and move directionally with cables at a similar rate (�0.3�m/s), and rapid patch movements are lost upon disruption ofcables by specific conditional mutants (159). Third, internal-ized patches/vesicles appear to fuse eventually with endosomalcompartments and shed their actin coats. The movement ofpatches on cables is consistent with an earlier live cell imagingstudy from fission yeast (289). Thus, slow patch movementsin the cell cortex (possibly to facilitate vesicle formationand scission) depend on an Arp2/3 complex-based actin poly-merization mechanism, whereas rapid inward movement ofpatches depends on a mechanism of transporting patches oncables. These findings raise further questions that remain un-resolved. What factors link patches to cables? How do cablesdeliver patches to endosomes, and are endosomes linked tocables? What signal triggers actin coat shedding at endosomes?

In animal cell endocytosis, short bursts of actin polymeriza-tion at the cell cortex (similar to actin patch assembly) areaccompanied by clathrin coat formation. Several yeast actinpatch components are physically linked to clathrin and itsadaptors, suggesting that clathrin may participate in patch dy-namics (21). However, until recently, the direct involvement ofclathrin in yeast endocytosis remained controversial, in part

because endocytic defects in clathrin mutants are not fullypenetrant, and in part because clathrin localization at corticalpatches had not been demonstrated (21). In addition, clathrin-coated pits had not been observed in EM studies on the yeastcell cortex (255, 314), but two recent studies have providedconvincing evidence that clathrin indeed localizes to earlypatches to facilitate endocytosis (174, 266). The localization ofclathrin at the cell cortex had been masked by abundant clath-rin staining on internal membranous structures. This technicallimitation was overcome using total internal reflection fluores-cence (TIRF) microscopy, which reduces fluorescent back-ground and permits focused observation of fluorescence at thecell cortex. Both studies used TIRF to show that clathrin isrecruited to early patches, possibly by Sla2, and then laterdisappears upon transition of patches to fast inward movement(174, 266). Further, it was shown that almost all clathrinpatches at the cortex subsequently recruit actin (i.e., becomeactin patches) and that clathrin mutants cause delays in earlystages of patch development. These studies have ended thedebate over whether clathrin functions in yeast endocytosisand raise exciting new questions, such as what signaling mech-anisms control recruitment and subsequent loss of clathrin atpatches. Another factor that plays a key role in endocytosis inanimal cells is dynamin, a GTPase that tubulates membranesand facilitates vesicle scission (298). Although multiple dy-

FIG. 2. Cortical patches are sites of endocytosis and actin assembly.(A) Colocalization of Abp1-GFP (an actin patch marker) and FM4-64(a membrane-binding dye/endosomal marker) at the yeast cell cortex.(B) Actin patches (Abp1-HcRed) undergo limited short-range move-ments while associated with the cell cortex and then transition to aphase of rapid movement upon internalization. The arrowhead tracksthe rapid movement of a single actin patch moving over time. Thepatch moves in a retrograde manner (bud to mother cell) on the endof an actin cable (marked by Abp140-GFP). The rate of fast patchmovement is approximately the same as the rate of cable movement(0.2 to 0.4 �m per s), suggesting that patches may be transportedpassively through attachment to cables. The molecular interactionslinking patches to cables are unknown. Bars, 2 �m. (Modified fromreference 159 by copyright permission of The Rockefeller UniversityPress.)



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namins are expressed in budding yeast, their involvement inendocytosis remains to be demonstrated (93).

Actin patch ultrastructure. Ultimately, to fully understandthe inner workings of actin patches, a detailed understandingof patch ultrastructure will be required at a resolution beyondthe limits of light microscopy. However, only a few studiesusing EM have been successful in elucidating details of patchultrastructure, due to the unique technical challenges pre-sented when analyzing actin in yeast cells: the presence of a cellwall, a concentrated and thus crowded cytoplasm, and a lowconcentration of actin filaments. Initial EM observations oftwo-dimensional thin slices revealed that actin patches associ-ate with finger-like invaginations (potential sites of endocyto-sis) at the plasma membrane (256) (Fig. 3A and B). Additionalwork on thin slices demonstrated limited colocalization of ac-tin with sites of receptor-mediated endocytosis (255) and dis-organized actin filament structures in endocytic mutants (340).

Using complementary EM techniques, a recent study byRodal et al. has provided the first three-dimensional informa-tion on patch ultrastructure (314). Intact samples of the yeastcell cortex, prepared using quick freezing and deep etching,revealed densely branched networks of actin filaments labeledwith antibodies to actin, Abp1, the Arp2/3 complex, and Crn1

(314) (Fig. 3C and D). The caveat to this methodology is thatit requires the use of spheroplasted cells. However, sphero-plasted cells maintain endocytosis (299), suggesting that thestructures preserved and visualized by EM represent func-tional actin patches. The structures are cone shaped, with theArp2/3 complex positioned at the apex, and there may be somerelationship between these actin mounds and the finger-likeinvaginations reported by thin sectioning (Fig. 3). In anotherstudy, Young et al. (423) used partial cell lysis to leach semi-intact actin patches from cells, purify them through multiplesteps, and image them by EM. The actin filaments in thesestructures form a branched network, and decoration with my-osin S1 fragment showed that the filaments have uniform po-larity. When combined with the in vivo observations of Rodalet al. (316) showing that the Arp2/3 complex (which binds tofilament pointed ends) is positioned at the apex of cone-shapedactin patches, these observations suggest that the majority ofactin filament barbed ends in actin patches may be orientedtowards the cell cortex. This assignment of filament polarityalso is supported by photobleaching experiments in sla2 mu-tants (173), which block patch internalization and lead to hy-perextended patch-like structures. A photobleached bandwithin this extended structure moves away from the cell cortex,suggesting that new monomers are incorporated on barbedends of filaments at the cortex.

Unified model for actin patch development and function. Inthis section, we provide a working model for actin patch de-velopment and endocytosis, based on the work described aboveand additional studies introduced below (Fig. 4).

(i) Early recruitment: nonmotile phase. The early recruit-ment process may begin with cytosolic regions of membranereceptors, possibly after being modified by kinases and ubiq-uitin ligases, associating with endocytic machinery (147, 156,323, 346, 362). Alternatively, some of the early endocytic ma-chinery may reside in patches at the cell cortex and recruit andconcentrate receptors. Supporting this second possibility, arecent study has described the existence of a static complex—termed the eisosome—that marks the future site of actin patchformation and contains the proteins Pil1, Lsp1, and Sur7 (390).It is important to note that actin patches also formed at sitesindependent of eisosomes, suggesting multiple mechanisms forspecifying sites of actin patch assembly. The initial endocyticmachinery recruited to patches includes clathrin and multiplescaffolds and clathrin adaptors: Yap1801 and Yap1802 (AP180homologues), Ent1 and Ent2 (epsin homologues), Ede1(Eps15R homologue), Scd5, Sla1, and Sla2 (6, 115, 145, 156,266). These proteins recruit additional early patch componentsthat promote actin assembly, such as Pan1, End3, and Las17, toform a complex network of interacting factors. Specifically, thenucleation-promoting factors (NPFs) (see below) Pan1 andLas17 may directly recruit and activate the Arp2/3 complex tonucleate actin assembly.

Of the many proteins involved in this early step of patchformation, two well-studied components that seem to play cen-tral roles are Sla1 and Sla2. Sla1 binds directly to receptorsto promote their internalization by a ubiquitin-independentmechanism (156). Sla1 also appears to be important for re-cruiting other factors to patches, as suggested by sla1� muta-tions causing delayed initiation of actin polymerization (i.e.,stalling in stage 1). Sla1 interacts with Pan1, End3 (364), Las17

FIG. 3. Electron micrographs of cortical actin patches. (A) Thinsectioning and electron microscopy reveal membrane invaginations atthe yeast cell cortex that are enriched for actin (immunolabeled with10-nm gold particles; arrow) and Abp1 (immunolabeled with 20-nmgold particles). (Reproduced from reference 256 by copyright permis-sion of the Rockefeller University Press.) (B) One model for actinfilament organization at cortical patches based on the microscopy inpanel A. (Reproduced from reference 256 by copyright permission ofthe Rockefeller University Press.) (C) Rapid-freeze deep-etch electronmicroscopy reveals actin filaments at the cell cortex organized intobranched, cone-shaped filamentous networks. Immunogold labeling(not shown) revealed that three known patch components (actin,Abp1, and Crn1) are distributed throughout these structures, whereasthe Arp2/3 complex is restricted to the peaks or apexes of the mounds.(Reprinted from reference 314 with permission of the publisher.)(D) Model for actin filament organization at patches based on theimages in panel C. The arrow indicates the direction of membraneinvagination during endocytosis.



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(213), and Sla2 (131). Sla1 has been shown to recruit Sla2 topatches (131). Sla2 localization to patches also requires Scd5,an essential protein that interacts with early patch componentsand regulates Glc7 phosphatase activity (145).

Sla2 is a multidomain protein that makes numerous contri-butions to patch development and function. First, Sla2 mayhelp recruit clathrin to patches, as suggested by the following:(i) Sla2 and clathrin colocalizing to early patches (266), (ii) the

mammalian homologue of Sla2 (Hip1R) colocalizing withclathrin in vivo and directly promoting assembly of clathrin invitro (94, 95), and (iii) two-hybrid interactions between clathrinand the Sla2 coiled-coil domain (144). Second, Sla2 may bindto and regulate Rvs167, the yeast homologue of amphiphysin.The BAR domains of amphiphysin/Rvs proteins promotemembrane curvature to facilitate vesicle budding in endocyto-sis (291). Third, Sla2 binds directly to filamentous actin via its

FIG. 4. Model for actin patch development. (Step 1) Receptors recruit early patch components, including clathrin, adaptors, and two NPFs(Pan1 and Las17), to the cell cortex to form a relatively immobile complex. (Step 2) Pan1 and Las17 recruit and activate the Arp2/3 complex tonucleate actin assembly, leading to slow patch movement at the cortex. During this stage, two additional NPFs are recruited, Abp1 and Myo3/Myo5.Abp1 in turn recruits the actin-regulating kinases Ark1 and Prk1, which phosphorylate Pan1 (and likely other patch proteins). Phosphorylationinhibits Pan1 activation of the Arp2/3 complex and triggers dissociation of the Pan1 complex (Pan1, End3, and Sla1). (Step 3) Pan1 phosphorylationand/or the activities of Rvs161, Rvs167, and the type I myosins Myo3 and Myo5 promote vesicle scission and internalization. The Arp2/3 complex,Abp1, Sac6, Cap1/Cap2, and likely other patch components maintain association with the internalized vesicle. Some early patch components areleft at the cell cortex, including Las17, Pan1, Sla1, Sla2, Myo3, and Myo5. (Step 4) Endocytic vesicles associate with actin cables by an unknownmechanism and move passively with actin cable retrograde flow (bud to mother). (Step 5) Cable-associated vesicles fuse with an endosomalcompartment concomitant with disassembly of the actin coat.



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carboxyl-terminal talin-like domain (239, 240). Although dele-tion of this domain from Sla2 causes no obvious defects inendocytosis (403, 420), this fragment of Sla2 is required for cellgrowth and endocytosis in the absence of specific clathrinadaptors, suggesting an important overlapping role in endocy-tosis (20). Thus, it will be interesting to learn if and how Sla2affects actin dynamics and organization. Fourth, through itsamino-terminal AP180 N-terminal homology domain, Sla2binds to phosphoinositide PI(4,5)P (PIP2) to facilitate the in-ternalization step of receptor-mediated endocytosis (360). Fi-nally, as Sla2 binds to Sla1 (18), which in turn binds two NPFs(Pan1 and Las17), it is possible that Sla2 also contributes to thespatial and temporal regulation of Arp2/3 complex activity.

(ii) Intermediate stage in patch development: slow motility.Transition to the second stage in patch maturation, whichlikely is coupled to membrane invagination, occurs when earlypatches (marked by Las17, Pan1, Sla1, and Sla2) are joined bythe actin nucleation machinery, consisting of the Arp2/3 com-plex and three new NPFs, Abp1, Myo3, and Myo5 (type Imyosins). Coincident with the detection of filamentous actin,patches begin to undergo slow, nondirectional movementswithin the plane of the cortex. These movements are sensitiveto Lat-A, suggesting that the transition to this intermediatestage is driven by actin polymerization (173). Using mutantsstalled at this stage of patch development, the measured ratesof actin polymerization were found to be 0.05 to 0.1 �m/s (173,174), markedly slower than subsequent cable-dependent rapidpatch movements (0.3 �m/s) (159).

(iii) Scission and rapid transport of patches. Transition tothe third phase of patch development correlates with the onsetof rapid, directional patch motility, in which patches/vesiclesmove inward from the cell cortex (173) and simultaneouslyshed many of their early components (e.g., Sla1, Sla2, Las17,Pan1, Myo3, and Myo5). The signals that trigger the transitionfrom slow-moving to fast-moving patches are unknown, butbased on several lines of evidence, Ark1/Prk1 kinases are im-plicated. These kinases associate directly with Abp1 and Sla2to localize to patches and directly phosphorylate Sla1 andPan1, and possibly other patch components (reviewed in ref-erence 355). Phosphorylation of Pan1 by Prk1 inhibits its abil-ity to bind actin and activate the Arp2/3 complex (371). Fur-ther, chemically induced loss of Ark1/Prk1 kinase activity invivo using analog-sensitive kinase mutants causes the rapid andArp2/3-dependent formation of actin clumps (340). Actinclumps consist of endocytic vesicles that have failed to matureproperly and are decorated with Abp1p, Sla2p, Pan1p, Sla1p,and Ent1p. Therefore, Ark1 and Prk1 may negatively regulatethe actin assembly-stimulating activity of early endocytic pro-teins and thereby play a critical role in directing vesicle inter-nalization and the onset of rapid patch movements.

In addition to regulation of Pan1 by Ark1 and Prk1 kinases,there are likely many other factors that contribute to theseevents. Las17 is shed prior to Pan1 (173), suggesting that Las17and Pan1 have different roles in vesicle internalization. Inaddition, mutations in Bbc1, End3, Sla1, and Sla2 (alone and incombinations) show formation of dramatic actin protrusions(174), suggesting that they may contribute to the down-regu-lation of actin assembly at this stage and/or help promotevesicle release. Consistent with this view, purified Sla1 andBbc1 directly bind to and inhibit the NPF activity of Las17, and

sla1� and sla2� cells show defects in patch ultrastructure byEM (314, 315). Deletion of SLA1 results in large, flattenedactin patches, and deletion of SLA2 results in large, raisedactin patches (314). Proper release of vesicles also requiresMyo3, Myo5, Rvs161, and Rvs167, which are implicated inpromoting membrane scission (172, 174). Rvs161 and Rvs167mutants in particular show a striking phenotype where inter-nalizing vesicles are retracted back to the cortex, suggesting adirect role in scission (174).

Once patches/vesicles leave the cell cortex, they move rap-idly inward along polarized actin cables, structures described indetail below. The retrograde (bud-to-mother) flow of cablesappears to “carry” patches to endosomal sorting compartments(159). The rate of patch transport is similar to the rate of cableflow, suggesting that transport is passive and must involve aphysical link between patches and cables (159). The only fac-tors known to associate with fast-moving patches are Abp1, theArp2/3 complex, Cap1/Cap2, and Sac6 (159, 173). Whetherother known F-actin-associated patch proteins and their li-gands (e.g., Cof1, Crn1, Srv2, Scp1, Twf1, and Abp140) arealso present in these patches remains to be determined. Ofthese factors, Abp140 and Sac6 are strong candidates for pro-viding linkage between patches and cables, since both areknown to decorate patches and cables (3, 17). Further, bothof these proteins can cross-link actin filaments in vitro (3, 17),an activity well tailored for bridging two filamentous actinstructures.

Finally, some late endosome movements have been reportedto require ongoing actin polymerization at the endosome sur-face. These movements were dependent on both the NPFactivity of Las17 and Lsb6 (a Las17-binding protein) (58, 59),and the authors suggested that Arp2/3-dependent actin assem-bly powers late endosome movements similar to the mecha-nism employed by Listeria (see above). However, one dilemmawith these observations is that Las17 and actin have not beendetected on late endosomes, which the authors suggest may bedue to their low abundance on such structures. Thus, it remainsopen whether these actin-dependent movements represent asecond, independent class of late endosomes, or instead thedefects in late endosome motility in las17 mutants arise fromaberrant assembly of endosomes at an earlier stage.

Actin Cables: Dynamic Tracks for Polarized Growth

Introduction. Unlike animal cells, which rely primarily onmicrotubule-based transport to establish and maintain cell po-larity (353), yeast cells use actin-based transport along cablesto direct polarized cell growth and to segregate organellesprior to cell division (45). Early in G1, unbudded cells detectcortical landmarks remaining from the previous cell division toselect a future bud site, and polarity factors (including thepolarity cap; see below) are recruited to this end of the cell.From this site, actin cable assembly is initiated, which leads toreorientation of actin cables and thus targeting of growth andsecretion to the future bud tip. Polarized growth towards thebud tip (or cap) continues through a medium-budded stageand depends on actin cables emanating from the bud tip andneck. These cables serve as polarized tracks for type V myosin-dependent delivery of cargos needed to build the daughter cell.

The first demonstration that actin cables are required for



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polarized cell growth was made possible by using a fast-actingtemperature-sensitive tpm1ts tpm2� mutant strain (306). In thiswork, Pruyne and coworkers showed a complete loss of cablesin tpm1ts tpm2� cells after a 1-minute incubation at the restric-tive temperature. Cells also rapidly lost the accumulation of asecretory marker (Sec4) and a class V myosin transporter(Myo2) at the bud tip. Remarkably, cables reassembledwithin 1 minute after their return to the permissive temper-ature, and polarity markers were restored at the bud tipshortly thereafter. Similar phenotypes were observed for thetemperature-sensitive myo2-66 allele, which carries a muta-tion in its motor domain (45, 171, 334). From an examina-tion of the literature, it is evident that mutations in mostfactors known to influence cable assembly and stability leadto cell polarity defects. These include tropomyosin, Sac6/fimbrin (4), capping protein (10), Srv2 (385), formins (97,99, 325), profilin (134, 410), and Bud6 (14).

Type V myosin motors (Myo2 and Myo4) transport diversecargos along actin cables to facilitate polarized growth. Post-Golgi secretory vesicles marked by GFP-Sec4 can be imaged inreal time moving directionally on cables towards the bud tip ina Myo2-dependent manner (335). These vesicles carry en-zymes that contribute to the production of new cell wall re-quired for bud growth and cell division (1, 343), includingglucan synthase, which localizes to actin patches (376). In ad-dition, organelles such as the vacuole, Golgi, nucleus, corticalendoplasmic reticulum, and peroxisomes (305, 421) are deliv-ered to the daughter cell in a cable- and Myo2-dependentmanner, and daughter-specific mRNAs (e.g., ASH1) are re-tained (and possibly transported) in the daughter cell in anactin cable- and Myo4-dependent manner (40, 72). Finally,mitochondria also appear to be transported along cables, andtheir anterograde movements occur through a myosin-inde-pendent mechanism that relies on Arp2/3 complex-based actinpolymerization, reminiscent of Listeria motility (104).

Mechanism of actin cable formation. Whereas patch forma-tion relies on actin nucleation by the Arp2/3 complex, cablesappear to assemble in an Arp2/3-independent manner (99,407). Until recent years, it had been speculated that cablesmight be assembled by a mechanism of filament capture andincorporation, similar to what was proposed for the assemblyof actomyosin rings and the bundled actin structures in Dro-sophila melanogaster bristles. However, recent studies showthat cables are assembled by the actin-nucleating activity offormins and profilin. Two key observations led to this discov-ery. First, it was shown that the formins Bni1 and Bnr1 areacutely required for cable assembly (99, 325), consistent withtheir localization to sites from which cables emanate (182, 284,304) (Fig. 5A). When conditional bni1ts bnr1� mutant cellswere shifted to the nonpermissive temperature, all visible actincables disappeared within 2 min (Fig. 5B), and cables reap-peared within 2 to 4 min upon return to the permissive tem-perature (99, 325). Second, biochemical analyses demonstratedthat purified carboxyl-terminal fragments of Bni1 directly nu-cleate actin assembly in vitro (303, 326). These papers inspiredstudies on formin activity in other species, and it is now evidentthat actin assembly-promoting activity is a universal propertyand function of all formins examined in animals, plants, andfungi (149, 194).

Despite these advances, key questions remain regarding ca-

ble formation. How are the potent actin nucleation activities ofBni1 and Bnr1 regulated? Studies in mammalian cells suggestthat formins are auto-inhibited until released by binding ofactivated Rho family GTPases to the formin Rho-binding do-main (389); however, it has not been determined if Bni1 and/orBnr1 are auto-inhibited. In yeast, there is evidence to suggestthat Cdc42 contributes to formin regulation during early budemergence and then the partially redundant Rho3 and Rho4regulate formins during most phases of cell growth, whereasRho1 may regulate formins in response to cell stresses (79).The specific molecular mechanism underlying this in vivo con-trol remains to be defined. Do Cdc42, Rho1, Rho3, and Rho4all act by binding to Bni1 and Bnr1 to relieve a proposedauto-inhibited state? Do they act specifically on Bni1 or Bnr1?Are other factors required for activating formins? How areBni1 and Bnr1 functions regulated by phosphorylation (122,235)? Once nucleated, how are actin filaments spatially orga-nized into cables? Is this mediated by the proteins that deco-

FIG. 5. Formins are required for actin cable assembly at the bud tipand neck. (A) Localization of GFP-labeled yeast formins Bni1 andBnr1 throughout the cell cycle. (Reprinted from reference 304 withpermission of the publisher.) (B) Formin function is required for theassembly and maintenance of actin cables. In a bnr1� background,point mutations in the FH2 domain of Bni1 render cells temperaturesensitive for growth and actin cable formation. Shifting mutants fromthe permissive temperature (23°C, left panel) to the nonpermissivetemperature (35°C, right panel) for 2 to 4 min results in complete lossof visible actin cables but does not affect actin patch distribution inbudded (b) or unbudded (u) cells. (Reprinted from reference 99 bypermission from Macmillan Publishers Ltd.)



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rate cables, such as Tpm1, Tpm2, Sac6, and Abp140 (17, 84,219, 306)?

The bud tip polarity cap. The term “polarity cap” refers toa group of interacting cellular factors that localize primarily tothe bud tip during bud emergence and growth and have geneticroles in directing polarized cell growth. Subsequently, many ofthe polarity cap components shift localization to the bud neck,just prior to cell division, where they appear to facilitate cyto-kinesis by mechanisms described in the next section. Somepolarity cap components help to promote cable assembly, asdiscussed below, while others regulate dynamic membranetrafficking events (exocytosis and endocytosis) and additionalaspects of cell polarity at the bud tip; for more details see otherrecent reviews (54, 138, 157, 302).

One functional complex within the polarity cap network hasbeen referred to as the polarisome. Its three components—Spa2, Pea2, and Bud6—comigrate as part of a 12S complex incell extracts fractionated by sedimentation velocity (345). Inaddition, the Rho family GTPase Cdc42 and two Cdc42 effec-tors, Bni1 and Gic2, appear to function intimately with thepolarisome, as supported by numerous two-hybrid and coim-munoprecipitation interactions with Spa2, Pea2, and Bud6 (97,113, 169). Dozens of other components now are routinelyreferred to as part of the polarity cap network based on theirphysical interactions and localization pattern to the bud tip.

The concept of a large macromolecular assemblage directingpolarized cell growth raises a number of interesting questions.First, what are all the components? To date, at least 60 pro-teins have been identified that localize to the bud tip, most ofwhich show physical and/or genetic interactions that suggesttheir involvement in regulating polarity (165). Undoubtedly,this list will continue to grow. Second, are there stable sub-complexes within the larger assemblage? As mentioned above,at least one stable complex has been identified, containingSpa2, Pea2, and Bud6, and there is evidence that this samecomplex may contain Gic2 (169) and possibly the Spa2-relatedprotein Sph1 (16, 319). Greater efforts are required now toisolate biochemically and define by mass spectrometry thecomponents of this and other bud tip complexes. Third, howare the interactions among complexes and components spa-tially and temporally regulated? As has proven to be the casefor other large biological machines (e.g., kinetochore, centro-some, and spliceosome), there are likely to be stable subcom-plexes as well as dynamically interacting components. Specificcomponents may change interactions in response to molecularsignals. Two kinases that regulate cell polarity are implicated inphosphorylating Bni1, a central figure in the polarisome be-cause of its role in nucleating actin assembly. Bni1 is phosphor-ylated directly by Fus3 kinase and is phosphorylated in a Ste20kinase-dependent manner in vivo (122, 235). The Bni1-inter-acting protein Bud6 is phosphorylated (251); further, it asso-ciates with Ste11 (a MEK kinase), which regulates polarity(251, 345). In addition, Bni1, Gic2, and the PAK-like kinasesSte20 and Cla4 are directly regulated by interactions with RhoGTPases (169, 170). These provide inspiring leads into themechanisms regulating polarity cap assembly and function butalso likely represent only a small percentage of the total sig-naling events involved.

A more daunting question, and one that will likely requirecombined efforts from many laboratories to answer, is what are

the activities of each component in the polarity cap network?Some of the relevant activities to address include the following:(i) maintaining association of other components at the cortex,(ii) providing signals to direct localized assembly of actin ca-bles, and (iii) receiving positive feedback signals that helpsustain polarity. Several polarity cap components (Bud6,Cdc42, Rho3, Rho4, and profilin) are thought to regulate Bni1directly to promote actin cable assembly (see below). In addi-tion, Spa2 directly interacts with Bni1 and Pea2 (113, 345), andSpa2 and Pea2 are required for Bni1 localization (284). Thebiochemical activity of Gic2 is unknown, but it binds to Cdc42(47, 60) and interacts with Bud6 in the yeast two-hybrid assay(169, 170), it contributes to localization of Bni1 and Bud6 inearly bud emergence (169), and gic1 gic2 mutants show defectsin establishment of polarity early in bud emergence (47, 60).Since Bud6 regulates Bni1 activity (253), it will be interestingto determine if Gic2 binds directly to Bud6 and whether Bud6-Gic2 interactions influence actin assembly.

With these and other data in mind, we have constructed aworking model for the polarity cap, focused on its role inpromoting actin cable assembly (Fig. 6). Future models willneed to incorporate mechanisms for secretory vesicle dockingand fusion, other membrane remodeling events, and cell wallsynthesis. In addition, a clear picture of polarity cap architec-ture and function eventually will require defining (i) the rela-tive abundances of each component, (ii) the stable proteincomplexes formed among components, (iii) the rules for hier-archical assembly and localization of components at the budtip, and (iv) functional interactions linking different nodes ofthe polarity network (e.g., bud site selection, Rho signaling,actin cable assembly, the exocyst complex, secretory apparatus,cell wall synthesis, RAM [regulation of Ace2p activity andcellular morphogenesis] signaling, and mitogen-activated pro-tein kinase signaling) (165). For instance, Msb3 and Msb4appear to coordinate multiple functions in polarized cellgrowth, including regulating the activity of the GTPase Sec4 inexocytosis, binding to the central polarity cap factor Spa2, andcontrolling Cdc42 activity (366).

Actin cable architecture. Cables must provide polarizedtracks for continuous myosin-dependent transport of cargosto the bud neck and tip. Therefore, early models suggestedthat cables might be stable structures, comprised of longbundled actin filaments, providing relatively stationary tracks formyosin movement. However, it is now evident that cablesare comprised of shorter filaments organized into bundlesof uniform polarity, with the majority of their fast-growing(barbed) ends oriented towards polarity sites. Evidence forthis organization includes the following: (i) a single fluores-cently labeled cable has variations in its width, suggestingthe thicker regions may be zones of intense filament overlap;(ii) cables are decorated with two known actin filament-bundling proteins, Sac6 and Abp140 (17, 84); (iii) myosinV-dependent transport is unidirectional towards polaritysites (335), and yeast type V myosins (Myo2 and Myo4) arebarbed end-directed motors (310); and finally, (iv) forminsassociate with the barbed ends of actin filaments and local-ize to the bud tip and neck. The precise arrangement ofactin filaments in S. cerevisiae cables remains uncertain,because cables traverse multiple planes within a cell andhave eluded description by thin-section EM and tomogra-



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phy. However, the recent use of permeabilized cells for EMin S. pombe has revealed that actin cables are comprised ofshort overlapping filaments with most of the barbed endsoriented towards the cell tips, consistent with facilitation ofbarbed end-directed myosin movement (175). Given all of

the points above, it seems likely that S. cerevisiae cables willshow a similar organization.

Gone in 60 seconds: actin cables are composed of shortfilaments. Intriguingly, actin cables are lost within 60 secondsof treating cells with the actin monomer-sequestering drug

FIG. 6. Models for regulation of actin cable assembly at the bud tip and neck. (A) Organization of formins (red), actin cables (gold), and septinstructures (blue) in a budded yeast cell. Formin proteins (Bni1 at the bud tip and Bnr1 at the neck) directly nucleate actin cable formation. Septinsserve as a diffusion barrier between bud and mother cell compartments and provide a scaffold for assembly of many bud neck components.(B) Interaction network at the bud neck: septins (blue); actin (tan); actin-binding proteins (red); all other factors (green). Solid lines representknown physical interactions; dashed lines denote septin-dependent localization. In G1/S, septins recruit other early components to the bud neck.Later in the cell cycle, just prior to cytokinesis, additional factors (late components) are recruited to the bud neck, including components of thebud tip “polarity cap” complex. (C) Model for regulation of actin cable assembly by polarity cap components at the bud tip and cable disassemblyby cofilin and Aip1. The polarity cap may be linked to the plasma membrane via components that directly associate with phospholipids, includingCdc42, Cdc24, and Cla4. The actin filaments in cables are nucleated by the formin Bni1, using profilin- and Bud6-bound actin subunits assubstrates. Bni1 activity may be regulated by interactions with Cdc42, Rho3, Rho4, Bud6, profilin, Spa2, and other factors (not pictured). Bni1associates tightly with the fast-growing (barbed) end of the actin filament it nucleates, facilitating insertional growth while protecting ends fromcapping protein (Cap1/2). Cables are likely comprised of short staggered filaments connected by actin cross-linking proteins, such as Sac6 andAbp140. It is not yet clear what regulates the lengths of actin filaments within a cable. Cables are stabilized along their sides by tropomyosin, whichcompetes with cofilin for binding F-actin. Genetic observations also suggest that Bni1 and Cap1/2 might cap the barbed ends of filaments in cables(see text). As rapidly as cables are formed, they must be disassembled, which recent evidence suggests is mediated by cofilin and Aip1.Barbed-end-directed type V myosins (Myo2 and Myo4) transport vesicles, organelles, and other cargos along cables to the bud tip.



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Lat-A (19). This finding demonstrates two important proper-ties. First, the filaments in cables likely undergo rapid turnover,as Lat-A sequesters monomers to inhibit new actin assemblybut is not reported to promote filament disassembly. Second,due to the rapid nature of cable loss, the filaments in cablesmust be relatively short. In fact, their approximate length canbe estimated from their disassembly rate using the followingsimple calculations. A typical large-budded yeast cell is �10�m long, with actin cables �4 �m long. Each actin subunitwithin a filament contributes 2.77 nm to filament length (155),so if this 4-�m cable is comprised of a single linear filament, itcontains �1,500 subunits. The barbed end of this filament maybe capped by formins (see below), but to make a more liberalestimate of filament length we will assume that both filamentends are uncapped. Using the dissociation rates for rabbitskeletal muscle ADP-actin (7.2 s�1 for the barbed end, 0.27 s�1

for the pointed end) and ATP-actin (1.4 s�1 for the barbedend, 0.8 s�1 for the pointed end) (293), a 4-�m filament (1,500subunits long) would lose about 450 subunits in 1 minute ifcomposed entirely of ADP-actin or 130 subunits for all ATP-actin. In either case, a 4-�m-long filament in a cable wouldmaintain most of its length after 1 minute of Lat-A treatment,instead of the observed complete loss of cable staining (19).Thus, an extremely conservative estimate of average filamentlength in cables is �1 �m. These estimations do not considerthe stabilizing and destabilizing effects of actin-binding pro-teins that decorate the sides of cables. Nonetheless, these es-timates for S. cerevisiae are remarkably consistent with themeasured lengths of filaments (0.4 to 0.5 �m) in S. pombecables as determined by EM (175).

This still leaves many open questions concerning cable reg-ulation. How is the length of filaments in cables controlled?The extent of filament elongation could be restricted by cap-ping proteins. While no conventional barbed or pointed endcapping proteins have been visualized on cables, it is possibletheir decoration is sparse (since they associate only with fila-ment ends) and thus has escaped detection. Cap1/Cap2 (yeastcapping protein), which associates tightly with filament barbedends to block both addition and dissociation of subunits, is astrong candidate to be on cables, because cap2� cells exhibitdramatically reduced cables (10). This phenotype is consistentwith Cap1/Cap2 having a role in stabilizing the filaments incables. Formins also bind tightly to barbed ends but permitfilament growth through a processive capping mechanismand protect growing ends from Cap1/Cap2 (253, 429). Thus,formins may initiate actin nucleation at the bud tip but remainassociated with filaments as they are incorporated into cables.Consistent with this model, actin cables undergo retrogradeflow away from the cortical sites of polarized growth (seebelow), and formin-GFP punctae have been visualized on mov-ing actin cables in S. cerevisiae and S. pombe (D. Pellman andF. Chang, personal communications). Thus, the capped stateof filaments in cables is an important issue to resolve and raiseseven more questions: are formins present on all or a subset offilaments in cables? Is Cap1/Cap2 present on some filaments?Do filaments initially have formins associated, but then Cap1/Cap2 displaces them? If so, what regulates these dynamics?Some of the issues may be resolved by emerging advances inlight microscopy, which may allow detection of a few (even

single) molecules decorating actin filament ends in vivo.Another possibility to consider is that filaments in cables

may be capped at their pointed ends. Tropomodulins performthis function in vertebrate muscle and nonmuscle cells, tightlycapping the pointed ends of tropomyosin-decorated filaments(108). Although obvious tropomodulin homologues have notbeen identified in the S. cerevisiae genome, functional homo-logues may exist, mutants of which likely would cause dimin-ished cables.

Actin cable dynamics and turnover. The observed rapidturnover of cables suggests that there may be active mecha-nisms required for disassembling the filaments comprising ca-bles. However, in wild-type cells, the primary filament disas-sembly factor, cofilin, is not detected on cables. Instead, cablesare decorated with tropomyosin, which stabilizes filaments andcompetes with cofilin for binding F-actin (30, 278). These ob-servations have left the high rate of cable turnover unex-plained; however, two studies provide insights into possibleturnover mechanisms. First, cofilin was shown to decorate ca-bles in aip1� cells (317). Second, cofilin and Aip1 were dem-onstrated to promote rapid cable turnover (276), with cof1-22and aip1� mutations causing �20- and 5-fold decreases in therate of cable turnover, respectively. This work suggests thatmost filaments in cables are decorated and stabilized by tropo-myosins Tpm1 and Tpm2, but subsets of filaments are stochas-tically and rapidly disassembled by the combined activities ofcofilin and Aip1 (see “Rapid Turnover of Actin Structures” formore details). This leads to a steady thinning or “pruning” ofcables along their lengths. Given the discovery of this functionfor cofilin and Aip1, other factors known to promote filamentdisassembly and turnover should be tested for their roles incable turnover (e.g., Srv2/CAP, profilin, and twinfilin).

Using Abp140-GFP as a marker, Yang and Pon have imagedactin cable dynamics in real time in live cells (419). Their workshows that for many cables, one end associates with the bud tipor bud neck while the other end moves away from these as-sembly sites (in the direction of the mother cell) at �0.3 �m/s.Some cables appear to assemble at the bud tip and extendthrough the bud neck into the mother compartment. Othercables appear to move along the cell cortex without having oneend attached to the bud tip or bud neck. It is not yet clearwhether these “free-roaming” cables have distinct functionsand whether they are initially assembled at the bud tip andneck and detach from those sites, or instead form independentof the polarity sites. The direction of cable flow (retrograde,away from the bud tip) is somewhat unexpected given thatcables serve as tracks for myosin V-based barbed end-directedtransport in the opposite direction (anterograde, toward thebud neck and tip). However, retrograde cable flow (0.3 �m/s)does not appear to pose a major difficulty to overcome, sincemyosin V-dependent anterograde transport is about 10 timesfaster—4.5 �m/s in vitro (310) and 3 �m/s in vivo (335).

What provides the force that drives cables away from thebud tip and neck and toward the mother cell? One possibleanswer is the force generated by actin polymerization. Asformins polymerize actin filaments at the bud tip and neck,these elongating filaments may be pushed into the mother celldue to insertional polymerization at their barbed ends cappedby Bni1 and Bnr1 (197, 251, 253, 429). If cable flow rate (0.3�m/s) were directly proportional to actin polymerization rate,



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this would require filament elongation rates of �100 sub-units/s. This would require 5- to 10-�M monomeric actin in thecytosol, which is unlikely to exist. However, interactions be-tween profilin, actin monomers, and the formins Bni1 andBnr1 may accelerate cable assembly rates to match rapid cableflow rates (see “Profilin-FH1 interactions: a throttle for fila-ment elongation” below).

In addition, cable flow rates are likely influenced by cellularmechanisms beyond actin polymerization, including a mecha-nism recently suggested to involve myosin. Domain and se-quence similarities among myosin heavy chains define myosinclasses, also called types (200, 400). S. cerevisiae expresses fivemyosins: two type V myosins (Myo2 and Myo4) that transportsecretory vesicles, organelles, and mRNAs to sites of polarizedgrowth (bud tip and neck); a single type II myosin (Myo1) thatpromotes formation and closure of the cytokinetic ring; andtwo type I myosins (Myo3 and Myo5) that promote actin as-sembly and endocytosis at cortical patches (22, 126, 305). Re-cent work suggests that Myo1 has an important role in con-trolling actin cable dynamics (T. Huckaba and L. Pon, personalcommunication). Myo1 first localizes to the incipient bud siteearly in G1 (33, 204, 362) and then is found at the bud neckfrom the earliest stages of bud formation. Although Myo1 isthought to function primarily in cytokinesis, much later in thecell cycle, Huckaba and colleagues observe reduced cable ve-locities in the absence of Myo1 motor activity. They proposethat Myo1, situated at the bud neck, uses its barbed end-directed motor to “pull” cables through the bud neck andfacilitate directional flow toward the mother cell. Thus, cablemovement may be regulated by actin polymerization, Myo1activity, and possibly other factors.

Yeast Cytokinesis: Two Complementary Mechanismsfor Cell Division

Introduction. Virtually all animals and fungi utilize an actin-based contractile ring to physically separate the two daughtercells during cytokinesis. Although a cytokinetic actin ring hadbeen observed in fission yeast as early as 1986 (231), the exis-tence of such a division mechanism in budding yeast was real-

ized only in recent years. Early work suggested an enrichmentof actin at the site of cell division in yeast (2, 183), but anactomyosin contractile ring was not observed until 1998 (34,217). In animal cells, myosin II controls contraction of the actincytokinetic ring (105), which constricts the cleavage furrow toa 1- to 2-�m-wide midbody zone (258, 308, 327). In S. cerevi-siae, the bud neck is less than 1 �m throughout the cell cycle;this led to the suggestion that constriction would not occur inyeast and, instead, that targeting of secretory vesicles for mem-brane and cell wall deposition to the bud neck would be suf-ficient to drive cell division (327). This view was reinforced bythe observation that the single budding yeast type II myosin,Myo1, localizes to the bud neck throughout the cell cycle(394), suggesting that Myo1 may have more of a structuralrole rather than a dynamic role at the bud neck. Further,since some yeast strains lacking MYO1 were shown to beviable and complete cytokinesis, its role in cytokinesis re-mained dubious (34, 318, 395). Then, in 1998, two seminalstudies imaged Myo1-GFP dynamics in live cells and pro-vided clear evidence that a myosin ring contracts specificallyduring cytokinesis (Fig. 7) (34, 217).

It is now evident that two complementary mechanisms con-tribute to yeast cytokinesis: first, contraction of an actomyosinring, and second, formation of a septum, achieved by deliveryof secretory vesicles to the bud neck to facilitate membranedeposition and cell wall synthesis. It remains unclear how thesetwo mechanisms are coordinated, as discussed below.

The position of actin ring assembly (and the future plane ofcell division) in budding yeast is dictated by the position of budemergence initiated in early G1, where the ring will be assem-bled at the neck of the newly growing bud (54, 302, 305). Thebud site is selected by cortical cues (bud site selection markers)remaining from the previous cell division. Early in G1, thesepositional cues trigger recruitment and activation of a cascadeof factors essential for assembling both the polarity cap at theincipient bud tip and components of the bud neck. At the topof this hierarchy of bud neck components are the septins,which form a ring structure discussed in detail below. Throughpoorly understood mechanisms, this scaffold in turn recruits

FIG. 7. Formation and contraction of an acto-myosin ring during yeast cytokinesis. (A) Prior to cytokinesis, actin filaments (rhodamine-phalloidin) and type II myosin (Myo1-GFP) assemble at the bud neck. (B) The actomyosin ring contracts, as demonstrated in consecutive framesof a Myo1-GFP time course at 1-minute intervals. Bar, 2 �m. (Reproduced from reference 34 by copyright permission of the Rockefeller UniversityPress.)



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Myo1, formins Bni1 and Bnr1, two myosin light chains (Mlc1and Mlc2), Hof1/Cyk2, IQGAP (Iqg1/Cyk1), and Cyk3 (22).All of these factors arrive at the bud neck before visible actinstaining with the exception of Cyk3, which appears duringanaphase.

During anaphase, two separate F-actin structures appear atthe bud neck, both of which contribute to cell division. First,actin cables become reorganized such that they are polarizedtowards the bud neck in both the mother and the daughter cellcompartments, directing all secretion to the division plane(305). This transition is marked by a dramatic shift of polaritymarkers (e.g., Cdc42, Bni1, Spa2, and Sec3) from the bud tip tothe bud neck, although the mechanism regulating relocation ofthese factors is unknown. These events lead to membrane andcell wall deposition at the bud neck to form the septum, dis-cussed in more detail below. Second, an F-actin ring forms,which constricts in a Myo1-dependent manner to help close theneck (34, 217). The actin filaments in the cables and the ringare highly dynamic, as both are sensitive to treatment of cellswith Lat-A (19, 34, 369).

These observations raise many important questions, in-cluding the following. What are the signals at anaphase thatinduce cable reorientation and actin ring assembly at theneck? What is the architecture of filaments in the ring?What are the signals that trigger ring contraction? How arethe activities of cables and the ring coordinated to facilitatecell division? Each of these issues is discussed in greaterdetail in the following sections.

Targeting secretion to the bud neck at cytokinesis. Prior toanaphase, polarized actin cables are assembled at two loca-tions, the bud tip (where Bni1 localizes) and the bud neck(where Bnr1 localizes). This produces two sets of cables thattogether target secretion to the bud tip. At anaphase, Bni1abruptly changes localization, joining Bnr1 at the bud neck(176, 284, 304). In a similar time frame, other components ofthe polarity cap, such as Spa2 and Bud6, change localizationfrom the bud tip to the bud neck (14, 345). These eventsredirect secretion to the bud neck, as indicated by relocation tothe neck of Myo2, Mlc1, and exocytosis markers such as Sec3and components of the exocyst complex (305). Targeted deliv-ery of new plasma membrane and cell wall materials providesadditional surface area required for cell division. Other keyfactors delivered to the neck include Chs2 and Chs3, the cat-alytic subunits of chitin synthase II and chitin synthase III.Chs2 contributes to formation of the primary septum, whileChs3 functions primarily in bud scar chitin synthesis; however,these two enzymes appear to share an essential role in septumformation (63, 343). chs2� and chs3� cells are viable but showdefects in septum formation and cell division, and chs2� chs3�double mutants are inviable and arrest at cytokinesis (343).This demonstrates an essential role for targeted septum depo-sition and cell wall synthesis in cytokinesis. Remarkably, thesesecretion-driven pathways are sufficient for completing cytoki-nesis in the absence of a contractile actin ring, albeit not asefficiently, suggesting that they make the dominant contribu-tion to cell division.

Formation and closure of the actomyosin ring. Actin fila-ments are among the final components to be assembled intothe contractile ring and do not become visible until late ana-phase. Early models, based on genetic evidence that the Arp2/3

complex is not required for actomyosin ring formation (408),suggested that preformed filaments may be recruited to thebud neck in order to form the ring (105). However, more-recent evidence suggests that ongoing actin assembly by Bni1and Bnr1 is required for ring formation and function. Whilethe actin ring forms in either bni1� or bnr1� single mutants(378), a temperature-sensitive formin mutant strain (bni1ts

bni1�) fails to assemble the ring (369). Other factors requiredfor ring assembly include profilin and tropomyosin (369). Thus,the machinery driving ring formation is highly similar to thatdriving cable assembly. In S. pombe, formins are essential foractin ring formation and cytokinesis (57), but the Arp2/3 com-plex also localizes to the actomyosin ring and makes a contri-bution to its formation (288), possibly by promoting endocy-tosis to facilitate septum formation. In contrast, actin ringformation in S. cerevisiae is thought to occur independently ofthe Arp2/3 complex, and to date the Arp2/3 complex has notbeen localized to the actin ring. Despite this prevailing model,two regulators of the Arp2/3 complex, Las17 and Vrp1, haveknown roles in cytokinesis (265, 367), and actin structures thatdepend on Arp2/3 complex activity have been observed at thebud neck (270), leaving open many possibilities for the mech-anism of their involvement.

Three questions regarding the behavior of the assembledactin ring point to important issues for future investigation.First, how are actin filaments in the ring organized? A require-ment for actin polymerization by formins suggests that fila-ments should be unbranched. Further, the classic “pursestring” model for ring contraction predicts a network of fila-ments with antiparallel arrangement to facilitate myosin-basedcontraction (109). Resolving this will require ultrastructuralanalysis of partially purified actomyosin rings and/or intactrings in cells. Second, what is the mechanism driving actin ringcontraction? One possibility is the motor activity of yeast typeII myosin (Myo1). Indeed, a point mutation impairing F-actinbinding of the Myo1 motor domain causes cytokinesis defects(T. Huckaba and L. Pon, personal communication). However,deletion of the entire motor domain of Myo1 causes no de-tectable defects in cytokinesis (224). While these data appearto be at odds, perhaps instead they suggest that the Myo1motor domain contributes to force production during cytoki-nesis but can be replaced by other cellular factors (possiblyother myosins) when deleted. Another possibility is that non-motor activities of Myo1 promote ring closure. Recent work inmammalian cells suggests that type II myosin promotes acto-myosin ring disassembly rather than contraction (133, 263).Myo1 may employ a similar mechanism to facilitate cytokine-sis, but this model awaits testing. It will also be important toidentify the signals that regulate Myo1 activity (and thus thetiming of ring contraction). Third, how are actin filaments inthe dynamic actomyosin ring disassembled? Myo1 may con-tribute to disassembly, as discussed above, but additional fac-tors are likely involved; one example is cofilin, which functionsat the cytokinetic ring in S. pombe (264). Clearly, we havemuch to learn about how the many ring components thatpromote actin filament assembly (e.g., formins, profilin), sta-bilization (e.g., tropomyosin, Sac6), and turnover (e.g., cofilin)are exquisitely regulated to control ring formation and con-traction.



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Role of yeast myosin light chains in cytokinesis. All myosinsare comprised of both heavy and light chains. The heavy chainsare large polypeptides (typically 100 to 200 kDa) with a motordomain, multiple IQ motifs, and variable tail regions. The lightchains are small polypeptides (�15 kDa) with a calmodulin-like fold that bind to IQ motifs in heavy chains to facilitatemotor activity. Further, calcium binding and/or phosphoryla-tion of light chains can regulate their association with heavychains, providing cells with a mechanism for controlling myosinactivity. S. cerevisiae expresses five myosin heavy chains (Myo1,Myo2, Myo3, Myo4, and Myo5) and three light chains (Mlc1,Mlc2, and the calmodulin Cmd1). Below, we focus on theirroles in cytokinesis through interactions with Myo1 and Myo2.

Like all type II myosins, Myo1 has an essential light chain(Mlc1) (227) and a regulatory light chain (Mlc2) (33). Differ-ences in the functions of the light chains are apparent fromtheir distinct patterns of localization and mutant phenotypes.Mlc2 colocalizes with Myo1 at the bud neck and requires Myo1for localization (227). In contrast, Mlc1 localizes to the budneck independently of Myo1 (342). Deletion of MLC2 haslittle effect on cell growth or cytokinesis but causes a slightdelay in Myo1 ring disassembly (227). In contrast, MLC1 isessential for cytokinesis and cell viability (33). Since myo1�cells are viable and can undergo cytokinesis, the lethality ofmlc1� suggests that it performs additional functions beyondregulating Myo1. This is further suggested by disruption ofMlc1-Myo1 interactions with specific point mutations inMlc1 or deletion of the IQ motifs in Myo1, both of whichcause only mild phenotypes (227).

What then is the primary Mlc1 function(s) in cytokinesis?Mlc1 binds to at least two other proteins at the bud neck thatpromote cytokinesis, Myo2 (358) and Iqg1/Cyk1 (IQGAP)(342). Its interaction with Myo2 targets vesicles to fill the budneck, a key step in formation of the septum (388). However,removal of all six IQ motifs from Myo2 (i.e., the myo2-�6IQmutant) does not affect growth or division rates (358), indicat-ing that the Mlc1-Myo2 interaction is not essential for motorfunction and likely serves primarily in a regulatory capacity.Mlc1 interactions with Iqg1 may have multiple roles. Iqg1requires Mlc1 for its localization to and function in assemblingthe cytokinetic ring (96, 341). Iqg1 also has been implicated inpromoting septum formation (280), which could require itsinteractions with Mlc1 and is consistent with the IQG1 genebeing essential (96). Thus, Mlc1 appears to coordinate bothactin-based pathways facilitating cytokinesis: (i) polarizedtransport of vesicles to form the septum and (ii) formation andclosure of the actomyosin ring. One key open question is howMlc1 localization is regulated, since there are no known li-gands of Mlc1 that it depends on for its localization to the ring.

Coordinating septum formation with actin ring contraction.Cytokinesis is driven by two parallel pathways (targeted vesicledelivery and ring assembly and closure) which are geneticallyseparable and are regulated by a large number of proteinsfound at the bud neck (121). The temporal sensitivity andcomplexity of these two pathways suggest that they must becoordinated; however, the mechanisms for this remain poorlyunderstood. One family of proteins that may be involved is theformins (Bni1 and Bnr1), as they directly nucleate the actinstructures required in both pathways: actin cables and rings.Their regulation—particularly by Rho-GTPases—may play a

key role in this process. Rho1, but not Cdc42, is required foractin ring formation, and the requirement for Rho1 is over-come by expression of a dominant-active Bni1 construct (369).This suggests that Rho1 functions upstream of Bni1 to stim-ulate actin ring assembly. Functions for other yeast RhoGTPases during cytokinesis remain untested.

Another strong candidate for coordinating septum forma-tion with contractile ring function is Hof1/Cyk2, a member ofthe cdc15/PSTPIP family of proteins (216). Deletion of HOF1/CYK2 causes temperature-sensitive growth and leads to defectsin both septum formation and ring contraction at the nonper-missive temperature (215, 378). Hof1 localizes to the bud neckindependent of Myo1 (215, 378) and contracts only partiallyduring cell division (unlike Myo1) (215, 378), and hof1 andmyo1 deletions are synthetic lethal (378). These observationsindicate that Hof1 and Myo1 perform distinct functions incytokinesis and suggest that Hof1 may function in septumformation. Hof1 physically interacts with Bnr1, suggesting itmay regulate cable assembly at the bud neck. Further, anothermember of the cdc15/PSTPIP protein family, S. pombeCdc15p, interacts with both the cytokinetic formin Cdc12p andthe Arp2/3-activating type I myosin (53). The genetic dataimplicating Hof1 in septum formation, along with physicalinteractions of Cdc15 and Hof1 with actin polymerization ma-chinery, together suggest that these proteins may link andcoordinate actomyosin ring formation/contraction with septumformation. These processes may be facilitated by the ability ofcdc15/PSTPIP proteins to associate with and deform mem-branes via their conserved FCH domains (166, 372).

A third factor to consider, Cyk3, localizes to the bud neck,shows genetic interactions similar to those of Hof1 (e.g., syn-thetic lethal with myo1) (193), and has been proposed to func-tion with Hof1 in septum formation (33). Overexpression ofeither CYK3 or HOF1 suppresses the lethality of iqg1� (193),suggesting that CYK3 and HOF1 may act downstream or areredundant with IQG1. Further, cyk3� and hof1� mutations aresynthetic lethal, suggesting that they may perform separatefunctions downstream of Iqg1 (193).

Septins. The molecular machinery that assembles the acto-myosin contractile ring and the septum must be targeted to thebud neck. The timing of arrival and loss of different compo-nents at the bud neck varies greatly throughout the cell cycle(121), and the mechanisms for recruitment and maintenance oflocalization of these proteins are just beginning to emerge.Some of the earliest proteins to arrive are the septins (includ-ing Cdc3, Cdc10, Cdc11, and Cdc12 in budding yeast), which inturn are required for localization of many other proteins (222).Septins are ubiquitous GTP-binding proteins that form stableheteromeric complexes; these units assemble tandemly into10-nm-wide filaments that can be organized further into poly-mer systems (106, 187). Septin filaments in cells were firstobserved by EM studies as striations at the bud neck thatdisperse in septin mutants (48, 49). Later, septins were shownby immunofluorescence to arrive at the cell cortex prior to budemergence (110, 184) and shortly thereafter reorganize into acollar that persists at the bud neck until after the completion ofcell division (167). Initial assembly of these septin structuresdepends on the activity of the GTPase Cdc42 (65, 119, 120,167) and two of its downstream effectors, Gic1 and Gic2 (167).



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Purified yeast septins assemble into filaments in vitro (111),and GTP binding is required for their polymerization and invivo functions (381).

Formation of the actomyosin ring depends on septin integ-rity (34, 217); however, only the Myo1 ring contracts (theseptin ring does not), indicating that they are separate struc-tures (110, 184, 222). In addition to positioning the actomyosinring, septins recruit enzymes, such as chitin synthase II, re-quired to form the septum (33). One model to explain thesecritical roles of septins is that they are near the top of a physicalinteraction network or hierarchy for assembly of componentsat the bud neck. However, potential physical interactions be-tween septins and the actomyosin ring machinery have re-mained elusive in yeast. In animal cells, the adaptor proteinanillin physically links actin and septin filaments in vitro (188).The fission yeast protein Mid1p has some sequence homologywith anillin and functions in placement of the contractile ringat the middle of the cell (56). However, Mid1p has not beenshown to bind to actin or septins; rather, Mid1p may interactwith type II myosin at the division site (254). No clear anillinhomologue exists in budding yeast, but Mid1 and anillin doshare limited sequence homology with the bud site selectionprotein Bud4 (29, 270, 328, 365). Further, Bud4 localizes to theneck (328) and binds Iqg1, consistent with a cytokinetic func-tion (280). So, Bud4 is a reasonable candidate for actin-septinlinkage functions. In addition, budding yeast Afr1 may facili-tate actin-septin interactions, as it associates with Iqg1 and theseptin protein Cdc12 (191, 279). However, the role of Afr1 maybe specialized for mating projection formation during the cel-lular response to pheromone (118).

Other recent work has defined a nonphysical anchoringmechanism by which septins maintain localization of someproteins at the bud neck. Prior to cytokinesis, the septin ringsplits into two separate rings (223), and this split ring structurecompartmentalizes and thereby concentrates (i.e., corrals) fac-tors such as Spa2 and Chs2 at the bud neck (77). In wild-typecells, Spa2 diffuses freely within this small zone between thesplit septin rings. However, in cdc12-6 mutants, Spa2 and Chs2rapidly diffuse away from the bud neck after cells are shifted tothe nonpermissive temperature and the split septin ring orga-nization is lost (77). Thus, septins may have dual roles inlocalizing cytokinetic machinery to the bud neck, serving pos-sibly as both physical scaffolds and diffusion barriers.

MECHANISMS OF ACTIN FILAMENT ASSEMBLY,ORGANIZATION, AND TURNOVER

Introduction

Below, we describe the properties and mechanisms of the S.cerevisiae actin-binding proteins known to affect actin dynamicsand/or organization, summarized in Table 1. In each case, weattempt to relate the activities of these proteins back to theircellular roles. Our discussion includes the type I myosins Myo3and Myo5, which help promote actin assembly, but not type IIand V myosins, which have been reviewed elsewhere in detail(44, 236, 305, 324, 373).

Biochemical properties of actin. Actin is a globular 42-kDaprotein that exists in dynamic equilibrium between two states,monomer and filamentous polymer. The double-helical fila-

ments assemble in a head-to-tail manner, imparting polarity tothe filament. The two ends of the filament are referred to asbarbed and pointed, owing to the arrowhead pattern observedupon decoration with the S1 fragment of myosin (296). Assem-bly of actin monomers into a filament involves an initial nu-cleation step, which is inherently slow due to instability of actindimers and trimers. However, once assembled, filaments arestable and undergo rapid polarized growth, with monomericATP-actin subunits preferentially adding to the barbed end(11.6 subunits �M�1 s�1) versus the pointed end (1.3 subunits�M�1 s�1) (293). Addition of an ATP-actin subunit to thebarbed end of a filament triggers hydrolysis of ATP bound tothat subunit (half-life [t1/2]), �5 seconds) (295), producingactin bound to ADP and Pi. In a subsequent (slower) step,actin subunits release Pi (t1/2, �5 min) to produce actin boundto ADP (295). Thus, filaments at steady state (undergoingaddition and loss at their ends) are a mosaic, enriched inATP-bound actin near barbed ends, ADP-Pi actin in the mid-dle, and ADP-actin at the pointed ends. ADP-actin subunitsdissociate from filament pointed ends, and the resulting ADP-actin monomer pool must undergo nucleotide exchange (ATPfor ADP) to “recharge” monomers for subsequent rounds ofbarbed end addition. This property of net subunit additionoccurring at barbed ends and net subunit dissociation occur-ring at pointed ends leads to steady-state cycling of subunitsthrough filaments, called treadmilling (Fig. 8) (268). Actinturnover refers to the collective dynamic events of actin sub-units treadmilling through filaments, dissociating from fila-ment ends and, as monomers, undergoing nucleotide exchangeto recycle them for new rounds of polymerization.

Each step in the process of actin filament assembly andturnover represents an opportunity for cellular control in re-modeling the actin cytoskeleton. All eukaryotic cells contain acore set of actin-binding proteins that regulate actin nucle-ation, barbed end addition, pointed end dissociation, filamentstability, and nucleotide exchange on monomers. For example,the Arp2/3 complex and formins bypass the inherently slowformation of actin dimers and trimers to nucleate actin poly-merization (194, 401). Filaments are also severed and rapidlydisassembled by factors such as actin-depolymerizing factor(ADF)/cofilin, Aip1, twinfilin, and Srv2/CAP (24, 268). Fila-ments can be stabilized by capping proteins and side-bindingproteins. Further, a number of actin monomer-binding pro-teins cooperate to replenish the assembly-competent pool ofactin monomers by catalyzing the conversion of ADP mono-mers dissociating from filament ends to ATP monomers. Theactivities and mechanisms of these factors are detailed in thissection, with emphasis on how these factors work togetherrather than alone to affect actin filament dynamics and orga-nization.

Assembly of Actin at Cortical Patches

The Arp2/3 complex. Actin patches arise at cortical siteswhere no preexisting filamentous actin can be detected and,thus, require de novo actin nucleation. Of the two known actinnucleators that are conserved from yeast to mammals (theArp2/3 complex and formins), only the Arp2/3 complex isfound at patches (248). Further, only the Arp2/3 complex has



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been implicated genetically in patch assembly (248, 407, 408).The Arp2/3 complex contains seven conserved subunits, in-

cluding two actin-related proteins (Arp2 and Arp3) and fiveunique proteins (Arc40/p40, Arc35/p35, Arc19/p20, Arc18/p21,

and Arc15/p15). The essential ARP2 gene was first identified inyeast based on sequence similarity to actin (337). Subsequentwork showed that arp2 temperature-sensitive mutants havedefects in endocytosis and actin organization (247, 248). The

TABLE 1. Biochemical activities and cellular localization of yeast actin-binding proteins

Name Homologue(s) Localization Classa Actin binding (Kd) Activities and functions Keyreferences

Abp1 mAbp1 Patches F Filament sides (�0.6 �M) Activates Arp2/3 complex; links Srv2 andArk1/Prk1 to patches

84, 127, 307

Abp140 Unknown Patches,cables

F Filament sides Bundles filaments; Abp140-GFP is used asa cable marker

17, 419

Aip1 Aip1 Patches F Caps barbed ends Promotes filament disassembly byenhancing cofilin severing and cappingcofilin-severed filaments

274, 275, 317

Arp2/3complex

Arp2/3 complex Patches F Filament sides (�2 �M)b;pointed ends (�20 nM)b

Nucleates filaments and promotes filamentbranching; requires activation by NPFs(Las17, Pan1, Myo3, Myo5, and Abp1)

127, 313, 406,408

Bni1 Formin Bud tip andneck

F Barbed ends (�7 nM)c Nucleates filaments; processively capsbarbed ends

303, 326, 414

Bnr1 Formin Bud neck F Barbed ends (�1 nM)c Nucleates and bundles filaments;processively caps barbed ends

251, 304

Cap1/Cap2 Capping protein Patches F Caps barbed ends (�10 nM) Caps barbed ends and stabilizes filaments 11, 415Crn1 Coronin Patches F Filament sides (�6 nM) Cross-links and bundles filaments; recruits

Arp2/3 to sides of filaments and restrictsnucleation activity to sides

128, 161

Iqg1/Cyk1 IQGAP Bud neck F Filament sides IQGAP bundles filaments and is regulatedby Cdc42 and calmodulin

26, 114, 341

Myo3 Myosin type I Patches F F-actin motor Activates Arp2/3 complex; may promotevesicle scission in endocytosis; regulatedby phosphorylation of TEDS motif

172, 210, 411

Myo5 Myosin type I Patches F F-actin motor Same as Myo3 See Myo3Myo1 Myosin type II Bud neck F F-actin motor Recruits filaments and promotes closure of

cytokinetic ring34, 217

Myo2 Myosin type V Bud tip andneck

F F-actin motor Nonprocessive motor; transports vesiclesand organelles to bud tip; facilitatesmitotic spindle orientation with Kar9and Bim1

171, 310, 421

Myo4 Myosin type V Bud tip andneck

F F-actin motor Transports daughter-specific mRNAs to budtip (e.g., Ash1)

40, 310

Pan1 Eps15 Patches F Filament sides (�1 �M) Activates Arp2/3 complex; binds filamentsvia atypical WH2 domain;phosphoregulated by Prk1 kinase

87, 371

Sac6 Fimbrin Patches,cables

F Filament sides (�0.5 �M)b Bundles and stabilizes filaments 4, 198

Scp1 Calponin/transgelin Patches F Filament sides (�1 �M) Bundles and stabilizes filaments 129, 405Sla2 HIP1R Patches F Filament sides Binds PIP2, clathrin, may bundle filaments;

HIP1R promotes clathrin assembly240, 360

Tpm1 Tropomyosin Cables F Filament sides (�2 �M) Stabilizes filaments; competes with cofilinfor filament binding

61, 81, 219,220

Tpm2 Tropomyosin Cables F Filament sides Same as Tpm1 61, 81Bud6/Aip3 Unknown Bud tip and

neckG ATP–G-actin (�0.75 �M)c Stimulates Bni1-mediated actin assembly;

promotes nucleotide exchange on actinmonomers

14, 253

Las17/Bee1 WASp Patches G Binds monomers Activates Arp2/3 complex; inhibited bySla1 and Bbc1

315, 406

Pfy1 Profilin Cytoplasmic G ATP–G-actin (�0.1 �M)b;ADP–G-actin (�0.5 �M)b

Promotes nucleotide exchange on actinmonomers; inhibits spontaneous actinassembly; restricts growth to barbed endsof filaments

88, 225, 410

Srv2 Cyclase-associatedprotein (CAP)

Patches G ATP–G-actin (�1.9 �M);ADP–G-actin (�20 nM)

Oligomerizes to form high-molecular-masscomplex (600 kDa) with actin monomers;blocks monomer addition to barbed ends;promotes conversion of cofilin-boundADP–G-actin to profilin-boundATP–G-actin

23, 214, 237,249

Cof1 Cofilin Patches F, G Filament sides (�1 �M);ATP–G-actin (�2 �M)b;ADP–G-actin (�0.1 �M)b

Severs/disassembles filaments; inhibitsnucleotide exchange on monomers

204, 205

Twf1 Twinfilin Patches F, G ADP–G-actin (�50 nM);filament sides

Sequesters monomers; severs filamentsat low pH; regulated by binding toCap1/Cap2 and PIP2

125, 252, 285

Vrp1 WASp-interactingprotein (WIP)

Patches F, G WIP binds filaments andmonomers

Binds Las17 and myosin type I; unknownbiochemical function

151, 179, 234,377, 382

a Proteins are listed alphabetically within three classes: binding to F-actin (F), binding to G-actin (G), and binding to both F-actin and G-actin (F, G).b Kd derived for nonyeast homologue.c Kd derived for yeast protein using nonyeast actin.



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other six subunits of the yeast Arp2/3 complex were identifiedin 1997 (407), and deletion of any subunit except ARC18causes severe growth defects or lethality (408). Further, theterminal phenotypes of these deletion strains show a severe orcomplete loss of cortical actin patch staining.

The Arp2/3 complex has been proposed to nucleate actinassembly by bringing together its Arp2 and Arp3 subunits tomimic the barbed end of a filament (294). This leaves thecomplex bound to the pointed end of the new filament withapparent nanomolar affinity (259). The complex also can bindto the sides of a preexisting (mother) actin filament and nu-cleate formation of a new (daughter) filament at a 70° angle,thus producing branched actin networks (9, 35, 150) (Fig. 9C).Alone, the Arp2/3 complex has minimal nucleation activity andrequires an activator or NPF for strong nucleation. The crystalstructure of the inactive bovine Arp2/3 complex reveals a largeseparation between the Arp2 and Arp3 subunits (313), andrecent studies using electron microscopy and fluorescence res-onance energy transfer analysis have demonstrated thatWASp/Las17 induces large structural rearrangements in thecomplex that bring Arp2 and Arp3 closer together to nucleateactin (Fig. 9) (124, 316). These observations in turn raise a newset of mechanistic questions. Which subunits facilitate Arp2/3complex binding to the sides of filaments? What are the preciserearrangements of subunits upon activation? Which subunits,and in what order, relay activation from WASp to the finalrepositioning of Arp2 and Arp3? Answers to these questionswill likely require mutagenesis of specific subunits involved,coupled with structural analyses of mutant and/or WASp-bound complexes.

Another fundamental property of Arp2/3 complex functionand regulation is binding and hydrolysis of ATP by the Arp2and Arp3 subunits. Arp2 and Arp3 both bind ATP (73, 209),but recent work suggests that only Arp2 hydrolyzes ATP (74,

208). Hydrolysis of ATP by Arp2 plays a critical role in WASp-induced conformational changes in the Arp2/3 complex (74,124). Further, hydrolysis by Arp2 and/or Arp3 may promotedebranching of filament networks by dissociating Arp2/3 com-plex from the pointed ends of filaments (208). Consistent withthe importance of ATP binding and hydrolysis for Arp2/3 com-plex activity in vitro, mutations in the nucleotide-bindingpocket of Arp2 and Arp3 severely impair actin patch dynamicsand endocytosis in vivo (233).

NPFs. The S. cerevisiae Arp2/3 complex is regulated in vivoby five NPFs: Las17, Myo3, Myo5, Pan1, and Abp1 (87, 98, 127,207, 406). Each NPF has an acidic motif that facilitates bindingto the Arp2/3 complex, possibly to the Arp3 and ARPC1/p40subunits (Fig. 10B) (181, 199, 286, 398). In addition, Las17binds to G-actin, whereas Myo3, Myo5, Pan1, and Abp1 bindto F-actin (Table 1). For Las17, Pan1, and Abp1, actin bindinghas been shown to be critical for their NPF activities (307, 315,371). Actin monomer-binding NPFs such as Las17 are thoughtto present their bound actin subunit to the Arp2 surface tofacilitate nucleation. Actin filament-binding NPFs are thoughtto recruit the Arp2/3 complex to the sides of preformed fila-ments to stimulate nucleation. Understanding the precise ac-tivation mechanism of each NPF represents an important chal-lenge for future research.

All five NPFs colocalize with the Arp2/3 complex, and theprimary function ascribed to each NPF is activation of Arp2/3-mediated actin assembly. However, each NPF arrives at adifferent stage of patch maturation. These and other geneticdata suggest that NPFs perform distinct yet genetically over-lapping functions in patch development (see below), possiblyacting sequentially on the Arp2/3 complex (173). Availablegenetic and biochemical data suggest that Las17 provides thestrongest NPF activity in forming patches, with Pan1 andMyo3/Myo5 being ancillary as NPFs. Further, the weaker NPF

FIG. 8. Assembly and turnover cycle of actin filaments. Actin filaments are polarized, with a fast-growing (barbed) end and a slow-growing(pointed) end. ATP-bound actin monomers (light gray) preferentially associate with the barbed end of the filament. ATP hydrolysis on an actinsubunit is triggered by its addition to the end of a filament. The rate of this reaction varies for actins from different species. After hydrolysis, ADPand Pi remain bound to actin (medium gray), followed shortly by the release of Pi to yield ADP-bound actin (dark gray). Thus, steady-state F-actinexists as a mosaic of actin subunits in different nucleotide-bound states. At steady state, the pointed ends of filaments undergo a net loss ofADP-actin subunits, generating a pool of ADP-actin monomers (G-actin). Following their dissociation, free monomers rapidly exchange nucleotide(ATP for ADP) to replenish an assembly-competent pool of ATP-bound G-actin. The actin monomer-binding protein profilin is one factor thatcan accelerate the rate of nucleotide exchange on G-actin. Profilin binds to the barbed end of an actin monomer (see cocrystal in right panel),sterically blocking the addition of profilin-bound actin monomers to the pointed ends of actin filaments, but not the barbed ends. Based on theseproperties and the high abundance of profilin in cells, it is hypothesized that most in vivo actin assembly occurs specifically at filament barbed ends.Upon addition of a profilin-bound actin subunit to a filament barbed end, profilin disassociates rapidly, leaving the barbed end free for additionof the next subunit. The coordinates for the profilin-actin complex were obtained from reference 336.



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FIG. 9. The Arp2/3 complex mechanism of actin assembly. (A) Projection structures of free and ligand-bound yeast Arp2/3 complexes (eachcomputed from 1,000 to 3,000 separate images). Electron microscopy and single-particle analyses were used to show that the Arp2/3 complex existsin an equilibrium among open, intermediate, and closed conformations (middle three panels), referring to the relative positions of the Arp2 andArp3 subunits within the complex. Binding of an inhibitor (Crn1/coronin) promotes the open conformation (left panel), while binding of an NPF(Las17/WASp) promotes the closed conformation (316). The open conformation is thought to represent the inactive state, while the closedconformation is “primed” for nucleation. (B) Model for structural rearrangements during activation of the Arp2/3 complex. Arrows indicateconformational changes leading to the next structure in the activation sequence. In the transition from the open to the intermediate conformation,Arp2 and Arp3 subunits move toward the center of the gap separating the two subunits. Next, there is a further closure of the gap between Arp2and Arp3 by a movement of Arp2 toward Arp3 and possibly a small movement of Arc18 toward the center of the gap, creating the closedconformation. WASp is shown binding to Arp2 and Arp3, and possibly also to p40, stabilizing the closed conformation. Coronin is shown bindingto p35, stabilizing the open conformation. (C) Model for nucleation of branched actin filaments by the Arp2/3 complex. Solid black arrows indicatephysical interactions; the dashed arrow depicts closure of the complex upon Las17 binding. The crystal structure of the bovine Arp2/3 complex(313) is docked into the three-dimensional EM reconstruction of the yeast Arp2/3 complex in its open inactive conformation (316). The complexassociates with the side of an existing (mother) filament and nucleates formation of a new (daughter) filament at a 70° angle (large red arrow).Nucleation is triggered by association of the Arp2/3 complex with Las17 and the side of the mother filament. These interactions promote majorstructural rearrangements within the Arp2/3 complex, bringing Arp2 and Arp3 closer together to nucleate an actin filament. Nucleation alsorequires Las17 binding to an actin monomer, presumably to provide the first subunit of the daughter filament. (All reprinted from reference 316by permission from Macmillan Publishers Ltd.)



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activity of Abp1 may promote patch transitions by competi-tively inhibiting stronger NPFs (71). Below, we discuss what isknown about the mechanism of each NPF and relate this totheir functions at patches.

(i) Las17/WASp. Las17 (also called Bee1) was the first NPFreported in yeast and was identified by its sequence homologywith mammalian WASp (213). Early studies showed thatlas17� cells have defects in patch organization, and a perme-abilized cell assay that measures incorporation of fluorescentactin was used to show that Las17 is required for actin assem-bly at cortical sites (213). Following demonstration that mam-malian WASp activates the Arp2/3 complex, a homologousWA fragment of Las17 (consisting of its WH2 [W] domain andacidic [A] motif) was shown to activate the yeast Arp2/3 com-plex (406). From this work, two key questions emerged regard-ing Las17 NPF function. First, how does Las17 activate theArp2/3 complex? Second, how is Las17 itself regulated? In-sights into both of these questions came after purification andbiochemical characterization of intact, full-length Las17 (315).It was revealed that full-length Las17 has an NPF activity farmore robust than that of its WA fragment, suggesting thatsequences N terminal to WA make major contributions toactivation of the Arp2/3 complex. This is also consistent withgenetic results showing that complete deletion of LAS17causes much more severe defects than deletion of the WAfragment alone (las17�WA) (406). Increased NPF activity forfull-length Las17 has been postulated to arise from N-terminalregions binding to F-actin and/or the Arp2/3 complex.

WASp family members (including SCAR/WAVE, which isnot conserved in yeast) can be regulated by one of two basic

mechanisms, auto-inhibition (in which the N and C terminiassociate directly to inhibit NPF activity) or transinhibition(mediated by direct interactions of additional proteins) (43). Inboth mechanisms, inhibition is relieved by signals from acti-vated Rho GTPases. For example, Cdc42 binds directly to theGTPase-binding domain (GBD) of N-WASp to relieve auto-inhibition (320), and Rac binds directly to one of the compo-nents of a proposed WAVE1 transinhibitory complex to re-lease active WAVE1 (89), although this mechanism remainscontroversial (reviewed in reference 359). It has been unclearhow yeast WASp (Las17) activity is controlled. Las17 lacks aGBD and fails to bind yeast Rho GTPases (213), and there areno obvious yeast homologues to the proteins comprising theWAVE transinhibitory complex. Biochemically, purified full-length Las17 is active (i.e., not auto-inhibited) and constitu-tively promotes Arp2/3 complex-mediated actin assembly(315). Two known Las17 ligands, Sla1 and Bbc1 (213, 370),directly inhibit but do not abolish Las17 activity (315). Thus, itseems likely that Las17 activity may be controlled by transin-hibition, similar to WAVE1. In addition, Las17 has many otherligands that may contribute to its transinhibition and release(126), including Vrp1 (WIP homologue) (98, 207, 229, 377),type I myosins (98, 207), and Rvs167p, Lsb1p, Lsb2p, Pin3/Lsb3p, Ysc84/Lsb4p, and Bzz1p (229, 370). A significant chal-lenge will be to determine how these different factors tempo-rally control Las17 activity at different stages of patch formation.

There also may be lessons to learn from studies in mammaliancells, where WIP binds stably to and inhibits N-WASp. This N-WASp–WIP complex is activated by Cdc42 specifically in thepresence of Toca-1 (homologous to Bzz1), which binds to Cdc42

FIG. 10. Yeast NPFs. (A) Schematics of each protein drawn to scale. Abbreviations: A, acidic; B, basic; CC, coiled-coil; EH, Eps15 homology;EVH1, Ena/VASP homology 1; IQ, IQ binding; LR, long repeat; PP or PPP, polyproline; SH3, Src homology 3; TH1/2, tail homology 1/2; WH1,WASp homology 1; WH2, WASp homology 2. The proposed actin-binding domain of each NPF is colored red; acidic domains are yellow. Las17binds to G-actin, while Abp1, Pan1, and Myo3/5 all bind specifically to F-actin. Pan1 contains many Ark1/Prk1 consensus phosphorylation sites inLR1 and LR2 (indicated by the circled P). (B) Sequence alignment of acidic domains from each yeast NPF, with the consensus sequence belowin bold.



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and N-WASp (151). All of the homologous proteins are ex-pressed in yeast, making it tempting to draw direct parallels be-tween these systems. However, Las17 and Bzz1 both lack Cdc42-binding domains. Further, bzz1� mutants show no obvious defectsin cell growth, actin organization, or endocytosis (356). Thus,Las17 regulation by these factors may be distinct in yeast and/orrequire additional proteins to achieve effects similar to animal cellinteractions of Cdc42-TOCA-WIP-WASp.

(ii) Pan1. Pan1 is recruited to early patches along withLas17, well before the arrival of the Arp2/3 complex and de-tection of filamentous actin. Pan1 could be recruited to patchesthrough its interactions with End3, Sla1, and/or clathrin adap-tors (363, 364, 402). pan1� mutations are inviable, but muta-tion of the Pan1 acidic domain (pan1-101, which abolishes NPFactivity) causes no obvious growth defects. This mutation doeshowever show synthetic growth defects when combined witharp2 alleles (71, 87, 402). Thus, it appears that Pan1 performsessential functions required for endocytosis and cell growthdistinct from its role as an NPF.

Pan1 NPF activity requires its A motif, which interacts di-rectly with the Arp2/3 complex, and an atypical WH2 domain,which binds directly to F-actin with low micromolar affinity(371). Through these interactions, Pan1 may recruit the Arp2/3complex to filament sides to stimulate nucleation and possiblypromote formation of branched actin networks, such as thoseseen at patches (314, 423). It is unclear when in patch devel-opment Pan1 functions as an NPF. However, its NPF activity isstrongly inhibited in vitro by Prk1 phosphorylation (371).These phosphorylation events could signal the observed loss ofPan1 from patches upon transition to rapid movements. Abp1may also play a role in these events, as it arrives just before thistransition and directly recruits Ark1/Prk1 kinases to patches(68, 102). Accordingly, abp1� cells show a delay in loss of Sla1from patches as they transition to rapid movement (174). Pre-sumably, Pan1 also remains on fast-moving patches in abp1�cells, since Sla1 and Pan1 form a complex that is disrupted byArk1/Prk1 phosphorylation (427).

(iii) Myo3 and Myo5. Just before the onset of rapid patchmovement, Las17 and Pan1 are joined by the genetically re-dundant type I myosins Myo3 and Myo5 (172, 174). Theseproteins are comprised of an N-terminal motor domain, alipid-binding TH1 domain, an F-actin-binding TH2 domain, anSH3 domain, and an Arp2/3 complex-binding A motif (Fig.10). Myo3 and Myo5, like Las17, are required for actin patchassembly in the permeabilized cell assay (see “Las17/WASp”above), and myo3 myo5 double mutants are synthetic lethalwith las17�WA mutants, suggesting that Myo3, Myo5, andLas17 are redundant as NPFs (98, 207). The NPF activities ofMyo3 and Myo5 have not yet been characterized biochemi-cally, but the TH2-SH3-A fragment of S. pombe myosin type Iactivates the Arp2/3 complex in vitro (210). This fragment bindswith very weak affinity to F-actin and does not bind G-actin,leaving the myosin I NPF mechanism unclear. One model is thatMyo3 and Myo5 NPF activities require interactions with Vrp1(WIP homologue), which binds to G-actin (15, 98, 116, 207, 411).Indeed, one recent study showed that purified S. pombe Vrp1enhances the NPF activity of the TH2-SH3-A fragment (352). Inaddition, it has been suggested that Myo3 and Myo5 function ina large complex that includes Las17 (98, 207).

Based on the late arrival of Myo3/Myo5 to patches and an

increased number of cortical invaginations seen by electronmicroscopy in myo3 myo5 mutants (172), it has been proposedthat Myo3 and Myo5 help promote the scission of endocyticvesicles from the membrane at the onset of rapid patch move-ment. Consistent with this possibility, fission yeast type I my-osin can alter the composition and organization of membranedomains (361). How the motor activity of myosin contributesto function awaits determination, but motor activity is requiredfor patch assembly in the permeabilized cell assay (207). Motoractivity could produce a specific arrangement of actin filamentsor, alternatively, transport a cargo such as the Arp2/3 complexto specific regions within a patch. Work on Acanthamoebacastellanii type I myosin (281) indicates a low duty ratio fortype I myosins, suggesting that processive movement by Myo3/Myo5 requires multiple motors on a cargo.

(iv) Abp1. Abp1 arrives at actin patches approximately whenMyo3 and Myo5 do, just prior to the onset of rapid patchmovement (173). However, once rapid movement commences,Myo3 and Myo5 are left behind at the cortex (172) while Abp1remains associated with patches throughout internalizationand transport on cables (159, 173). abp1� mutants fail to shedSla1 and possibly other early patch components during thetransition to fast patch movement, suggesting that Abp1 hasa role in disassembly of the endocytic structures (174). Con-sistent with this model, Abp1 recruits Ark1 and Prk1 kinasesto patches (68, 102); these kinases phosphorylate Pan1 todisrupt the Sla1-Pan1-End3 complex (427). This model alsois consistent with Abp1’s role in recruiting to patches Sjl1(synaptojanin homologue), which is implicated in endocyticcoat disassembly (357).

The biochemical properties of Abp1 provide a plausibleexplanation for these functions. Abp1 is comprised of an N-terminal ADF/cofilin homology (ADFH) actin-binding domain(ABD), two A motifs, a polyproline region, and a C-terminalSH3 domain (Fig. 10). The NPF activity of Abp1 requires notonly its A motifs but also its ADFH domain, through which itbinds directly to F-actin with submicromolar affinity (127).These properties allow Abp1 to recruit the Arp2/3 complex tothe sides of preexisting filaments, which leads to its modestNPF activity. The abilities of Abp1 to bind both F-actin and theArp2/3 complex are required in vitro for Abp1 NPF activityand in vivo for its shared genetic functions with Sac6 (307).Similar to its mammalian relative cortactin (375, 399), Abp1may displace Las17/WASp from the Arp2/3 complex and sta-bilize filament branches, but this remains to be tested.

What is the functional relationship of Abp1 with other NPFs?With its two A motifs, Abp1 binds to the Arp2/3 complex withhigh affinity, yet has notably weak NPF activity compared tofull-length Las17 and Pan1 (127). These properties are well-suitedfor Abp1 acting as a competitive antagonist, which is supported byAbp1 attenuation of Las17 NPF activity in vitro and by geneticinteractions of abp1� with other NPF mutants (71). It is alsoconsistent with the appearance of Abp1 at patches after otherNPFs (173). Thus, sequential interactions of NPFs with theArp2/3 complex may regulate the transition from slow to fastpatch movement.

In addition to its NPF role, Abp1 recruits Srv2/CAP topatches by physically linking it to F-actin (23, 82, 214). As such,Abp1 may facilitate rapid turnover of F-actin, which is re-



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quired for slow patch movements prior to the onset of rapidpatch movement (173).

Coronin: spatial inhibitor of Arp2/3 complex activity. A novelform of Arp2/3 complex regulation, distinct from NPF activity, ismediated by the yeast homologue of coronin Crn1, which local-izes to actin patches (128, 142). Crn1 was isolated initially fromcrude extracts on microtubule affinity columns as a factor coelut-ing with actin (128). Recombinant Crn1 binds with high affinity toF-actin and cross-links F-actin to microtubules. F-actin binding ismediated by an amino-terminal WD repeat �-propeller structure,although the actin-binding surface has not been identified. Thisdomain is followed by a 200-residue-long “unique” region ofunknown structure that mediates microtubule binding and is notconserved in other species. At the C terminus is a coiled-coildomain that mediates Crn1 oligomerization and is required forF-actin cross-linking and bundling. Independent of F-actin cross-linking, Crn1 also modestly promotes actin assembly; however,the mechanism underlying this activity remains unclear. The�-propeller and coiled-coil domains are conserved in coroninsfrom diverse species, as are the abilities to bind and cross-linkF-actin (75).

The coiled-coil domain of Crn1 also binds directly to theArp2/3 complex, through interactions with the ARPC2/p35 sub-unit. This interaction directly inhibits actin nucleation by theArp2/3 complex (161), arising from direct Crn1-Arp2/3 interac-tions (NPF independent) that stabilize the Arp2/3 complex in itsopen (inactive) conformation (316). In electron micrographs ofthe Crn1-bound Arp2/3 complex, the globular �-propeller do-main rests �20 Å from the ARPC2/p35 subunit (Fig. 9A). Thecoiled-coil and unique regions mediating this interaction are notvisible, presumably because they lack substantial mass and/ortertiary structure. Since the mammalian coronin Cor1 also coim-munoprecipitates with the Arp2/3 complex, this regulatory func-tion of coronin may be widely conserved (50).

What is the cellular function of Arp2/3 complex inhibition byCrn1? While the coiled-coil domain of Crn1 mediates inhibi-tion, the presence of preformed actin filaments relieves inhi-bition and thereby permits Arp2/3 nucleation activity (161).Thus, Crn1 appears to spatially restrict Arp2/3 activity to thesides of preexisting filaments by suppressing de novo nucle-ation in the absence of filaments. As such, Crn1 may promoteformation of branched filament networks, a model that can betested by characterizing Crn1 effects on filament branching bythe Arp2/3 complex.

Crn1 localization to cortical patches requires both its �-pro-peller and coiled-coil domains, suggesting requirements forboth F-actin binding and coiled-coil-mediated functions (oli-gomerization and/or Arp2/3 complex binding). Although thetiming of its arrival to patches has not been reported, it may besimilar to that of other F-actin-binding proteins, Abp1, Sac6,and Cap1/Cap2, arriving late and persisting on patches throughrapid movements. Surprisingly for a factor with such potentinhibitory activity on the Arp2/3 complex, deletion of CRN1does not result in detectable growth or morphological defects(128). However, a physiological role for Crn1 in regulating theArp2/3 complex is supported by synthetic defects betweencrn1� and arp2-21 mutations (161). Further, crn1� has syn-thetic defects in cell growth and actin organization with cof1-22and act1-159 (mutations that slow actin turnover), suggesting arole in promoting actin turnover. However, the mechanistic

basis of such a function remains undetermined.Several aspects of Crn1 function that remain to be resolved

include the following. How does Crn1 regulation of Arp2/3complex activity contribute to patch maturation and endocy-tosis? Mechanistically, how does the presence of F-actin over-ride Crn1 inhibition of the Arp2/3 complex? Does Crn1 regu-late actin turnover in vivo with cofilin? Does Crn1 interact withmicrotubules in vivo, and how might this be coordinated withits actin functions?

Open questions about the Arp2/3 complex. In addition todirect inhibition of the Arp2/3 complex by Crn1, binding ofcertain proteins (e.g., tropomyosin) to actin filaments prohibitsArp2/3 complex association and thus indirectly inhibits theArp2/3 complex (37). This may explain why the Arp2/3 com-plex is not detected on actin cables, which are decorated byTpm1 and Tpm2. It also raises an important mechanistic ques-tion: how do cables and not patches selectively become deco-rated with Tpm1 and Tpm2? Another factor that may regulateArp2/3 complex function is Cmd1 (calmodulin), which inter-acts both genetically and biochemically with the ARPC2/p35subunit (330). There are likely many other cellular factors thatmust be coordinated with NPFs and Crn1 to control Arp2/3complex function.

There are also many open questions regarding Arp2/3 com-plex cellular function. First, what is the specific role of Arp2/3complex-based actin polymerization in patch maturation andendocytosis? The Arp2/3 complex is required for actin assem-bly, and actin assembly is required for endocytosis, but whatfunction does this provide? Actin polymerization could pro-vide force generation to drive vesicle scission; alternatively, itcould provide a dynamic scaffold to organize endocytic factorsthat drive membrane invagination and scission (93). There arealso non-patch/endocytosis functions of the Arp2/3 complex toconsider. For instance, the involvement of the Arp2/3 complexin cable and cytokinetic ring assembly has not been ruled out.Although studies on arp2, arp3, and arc35 mutants have notindicated a role in cable assembly (99, 407, 408), given thephysical links between patches and cables (159, 177) and mul-tiple reports suggesting interactions between the Arp2/3 com-plex and formins (69, 152), this question deserves further in-quiry. Further, while one study in budding yeast suggests thatthe actin ring is assembled by formins and not the Arp2/3complex (369), a separate study in fission yeast reports a rolefor the Arp2/3 complex in ring assembly (288). Thus, addi-tional investigations are needed to resolve this potentialArp2/3 complex function at cytokinetic rings. Finally, separatefrom assembly of the three major actin structures—patches,cables, and rings—Arp2/3 complex function contributes toorganelle inheritance, nuclear export, and mitotic spindle ori-entation (41, 330, 331, 416). Understanding the mechanisticbasis of these functions could broaden the range of action forthe Arp2/3 complex.

Assembly of Actin Cables

Formins Bni1 and Bnr1. Actin cables are essential for po-larized cell growth, but until recently the mechanism of actincable assembly was unknown. Early cell biology studies sug-gested a role for the two budding yeast formins Bni1 and Bnr1in cable formation (97, 164, 176, 182, 284), but it was hypoth-



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esized they had indirect roles in this process, serving as mo-lecular scaffolds. Further, technical obstacles in purifying theselarge proteins (�200 kDa) delayed characterizing their effectson actin in vitro. Similar to all formins of the Diaphanous-related formin (DRF) family, Bni1 and Bnr1 are comprised ofwell-defined domains that differentially contribute to theirmechanism and regulation (Fig. 11A). These include theformin homology 2 (FH2) domain, FH1 domain, GBD, Diaph-anous inhibitory domain (DID), and Diaphanous autoregula-tory domain (DAD) (100, 389). A critical breakthrough inunderstanding the role of formins in regulating the actincytoskeleton came when two studies showed that the FH2domain of Bni1 is critical for cable formation (99, 325).

Biochemical studies closely followed, demonstrating that puri-fied FH2-containing fragments of Bni1 directly nucleate actinassembly (303, 326). Similar actin-nucleating activities havesince been reported for formins from a wide variety of animals,fungi, and plants, making it clear that actin assembly is aconserved function of formins (149).

Budding yeast expresses two formins, Bni1 and Bnr1 (164),which partially overlap in their localization to the bud tip andbud neck (Fig. 5A). Bni1 and Bnr1 also have distinct andoverlapping functions in the assembly of cables and the acto-myosin ring (99, 304, 325, 369). A comparison of their bio-chemical activities reveals differences: Bnr1 specifically bun-dles actin filaments and has 10- to 15-fold more potent actin

FIG. 11. Formin mechanism of actin assembly. (A) Domain organization of budding yeast formins Bni1 and Bnr1. SBD, Spa2-binding domain.(B) Crystal structure of the Bni1 FH2 dimer depicted by surface rendering, with a solid line separating the two functional halves (hemi-dimers).Residues are colored according to the degree of evolutionary conservation, with highly conserved residues in red and nonconserved residues in tealblue. Circled regions denote patches of conserved surface residues involved in actin binding, identified by mutational studies (348, 414) (Reprintedfrom reference 414 with permission from Elsevier.) (C) Cocrystal structure of the Bni1 FH2 domain associated with tetramethylrhodamine-actin.Each hemi-dimer interacts with two actin subunits in the polymer. (Reprinted from reference 282 by permission from Macmillan Publishers Ltd.)(D) Model for processive capping by the FH2 dimer, which maintains association with the rapidly growing barbed end of the actin filament.Alternating regions of the FH2 are displaced rapidly in response to subunit addition to allow insertion of the next subunit. Importantly, the precisebiophysical mechanism of formin processive capping remains unclear. (E) While the formin caps the barbed end of the filament it has nucleated,the Arp2/3 complex caps the opposite (pointed) end of the filament, leaving the barbed end free. Thus, filaments nucleated specifically by forminsare protected from early termination of elongation by capping protein. Directions of new filament growth are noted by red arrows.



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assembly than that of Bni1 in vitro (251). Bni1 and Bnr1 alsoappear to be regulated by distinct mechanisms (see below).How these differences in activity and regulation tailor Bni1 andBnr1 to their respective cellular functions remains to bedetermined.

What is the formin mechanism of actin assembly? Whereasthe Arp2/3 complex bypasses kinetic barriers to actin assemblyby mimicking the barbed end of an actin filament, formins usea different mechanism. Kinetic studies suggest that the FH2domain promotes filament assembly by stabilizing polymeriza-tion intermediates, actin dimers and/or trimers, which are oth-erwise extremely short-lived species (300). This is facilitated byformins having very high affinity for F-actin. Once nucleated,formins remain bound to the rapidly growing barbed end of thefilament while guiding insertional addition of actin subunits.This activity is referred to as processive capping. The firstindications of this activity came from the demonstration thatBni1 FH2 localizes to filament barbed ends by immunoelec-tron microscopy (303) and does not completely inhibit barbedend growth (300, 303). Two subsequent studies showed thatBni1 FH2 protects elongating barbed ends of filaments fromthe inhibitory effects of conventional capping proteins (253,429). This process, which involves a strong interaction betweenFH2 and filament barbed ends (Kd � 20 nM) (253, 300, 303),has now been visualized in real time using TIRF microscopy ofindividual FH2-capped actin filaments (197) and drives rapidbarbed end filament growth at various rates (194, 197, 300, 303,429).

While a thorough mechanistic understanding of processivecapping is still far off, critical insights have been gained byrecent structural analyses. First, the FH2 dimerizes, and pointmutations disrupting dimerization abolish actin assembly(253). This suggested that a two-headed actin-binding struc-ture was required, opening the possibility of alternating actin-binding contacts to allow insertional growth and persistentassociation with the filament barbed end. Shortly thereafter,the crystal structure of the Bni1 FH2 domain was solved, whichrevealed a flexibly tethered dimer in which each FH2 moleculehas two actin-binding sites (Fig. 11B) (414). Sites of contactbetween FH2 and actin have since been defined at an atomiclevel by a cocrystal structure of the Bni1 FH2 domain bound totetramethylrhodamine-labeled actin (282) (Fig. 11C).

Based on these findings, several different models for proces-sive capping have been proposed. First, the stair-steppingmodel envisions two flexibly tethered actin-binding heads al-ternating interactions with the growing barbed end of the fil-ament (414). Anticooperative conformational changes in FH2drive alternating contacts to allow subunit addition (414). Acaveat to this mechanism, however, is that it requires rapidrotation of the FH2 dimer to account for a helical twist of theactin filament, and experimental evidence suggests that such arotation may not occur (197). To resolve this paradox, a secondmodel suggests that the unidirectional twist induced by multi-ple stair steps may be relieved by a periodic “backward” step,in which the FH2 twists in the opposite direction as the fila-ment (344). A third model proposes that the FH2 dimer mayact like a shaft in a bearing, encircling the growing actin fila-ment (197). However, the molecular contacts that maintainpersistent FH2 association with the growing barbed end in thismodel are not clear. Finally, a fourth model suggests that the

FH2 may adopt an end-side-binding mechanism, whereby itassociates with the side of barbed ends to allow new subunitaddition while prohibiting binding of capping proteins (137).Experimental evidence has not yet ruled out any of thesemechanisms and, in fact, they may be more similar mechanis-tically than they are distinct. Resolving the mechanism mayrequire the use of high-resolution electron microscopy tech-niques to image FH2 dimers bound to the barbed ends offilaments. Similar techniques have been used to define confor-mational changes in the Arp2/3 complex upon WASp activa-tion and at filament branches (91, 316) and to image proteincomplexes at microtubule ends (422).

While recent efforts have focused on defining the forminmechanism of actin assembly, many critical questions remainopen surrounding formin regulation. Like WASp and theArp2/3 complex, the potent activities of formins are likelysubject to tight spatial and temporal regulation. Below, wediscuss what is known about the regulation of Bni1 and Bnr1 bytheir known ligands, which include Rho GTPases, profilin,Bud6/Aip3, Tef1 (eEF1� homologue), and Spa2.

Profilin-FH1 interactions: a throttle for filament elongation.Profilin is an abundant actin monomer-binding protein foundin all eukaryotic cells examined. Further, profilin-bound actinmonomers are considered the primary physiological substrateavailable for actin assembly in vivo. Profilin has two separatebinding surfaces, one that binds actin monomers and anotherthat binds polyproline motifs, including those found in FH1domains (225, 410). Indeed, early studies identified profilin asa ligand of Bni1 and Bnr1 (97, 164). Profilin binding blocksactin monomer addition to the pointed ends of filaments butfreely allows addition to barbed ends (297), consistent with theprofilin-actin cocrystal structure (Fig. 8) (336). These proper-ties of profilin are fundamental to how it contributes to theformin mechanism of actin assembly.

Importantly, the Arp2/3 complex nucleates actin filamentswith free barbed ends and, thus, Arp2/3-nucleated filamentselongate at similar rates in the presence and absence of profilin(315). In contrast, formins remain tightly associated with thebarbed ends of filaments they nucleate and sterically inhibitcapping protein association, raising an important mechanisticquestion: can profilin-bound actin monomers be added effi-ciently to the barbed ends of formin-capped filaments? Thefirst insights came from Sagot et al. (326), who found that Bni1FH1-FH2 assembles actin filaments equally well from a pool offree actin monomers or from profilin-bound actin monomers.Further, FH1-profilin binding is required for formins to assem-ble profilin-bound actin monomers (300, 326). Thus, FH2alone promotes filament assembly from free actin monomers,but FH1 is required for filament assembly from profilin-boundactin monomers. This emphasizes the importance of the FH1domain for formin function in cells, where profilin-bound actinmonomers are the primary substrate available for filamentassembly. As such, FH1 is required for formin-mediated actinassembly in vivo, which is supported by many studies (99, 148,196, 253, 303, 325, 326, 392).

More-recent biochemical studies have revealed exciting ef-fects on actin assembly resulting from profilin-FH1 interac-tions with profound implications for formin function in vivo.By studying an isoform of mammalian formin mDia1 withextended polyproline tracts in its FH1, Romero and coworkers



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demonstrated a formin/profilin-dependent increase in rate ofbarbed end elongation—many times that of the diffusion-lim-ited rate of free barbed end growth (321). Prior to this work, itwas a widely held view that actin polymerization could notexceed diffusion-limited rates of assembly (�11 subunits persecond per micromolar). Thus, in a solution of 10 �M G-actin,filaments would elongate at �110 subunits per second. Thework of Romero et al. showed that filaments capped by form-ins can elongate at least fivefold faster specifically when pro-filin is present. A more recent study now has used TIRF mi-croscopy to elegantly demonstrate these effects in real time forindividual actin filaments capped by a variety of formins, in-cluding Bni1 (194). Further, they show that the magnitude ofthe increase in filament elongation rate correlates with thenumber of profilin-binding sites in FH1. The largest increasemeasured (�5-fold) occurs on filaments capped by mDia1isoforms with 5 to 15 profilin-binding sites. By comparison,Bni1 has three predicted profilin-binding sites and exhibits an�2-fold increase in rate of elongation (over the rate of elon-gation at free barbed ends). Bnr1 has not yet been examined,but it has two predicted profilin-binding sites, suggesting itlikely increases the rate of elongation by no more than twofold.

These results have important implications for formin func-tion in vivo. The yeast actin cable flow rate is �0.3 �m/s (419).If cable flow is driven primarily by actin polymerization, assuggested, this corresponds to addition of �100 subunits/s atfilament ends, requiring a cellular concentration of �9 �MG-actin. However, the total concentration of actin in yeast cellsis estimated to be 12 �M (A. Goodman and B. Goode, unpub-lished data), the majority of which is thought to be in filamen-tous form (178). Thus, the G-actin concentration in yeast cy-tosol should be well below 5 �M. These discrepancies canbegin to be reconciled by considering the formin/profilin-de-pendent increase in elongation rates. Further, this may explainwhy measured rates of actin assembly at patches (0.05 to 0.1�m/s) are markedly lower than cable flow rates (173, 174).

Although it is now clear that the FH1 domain serves as athrottle controlling the speed of elongation at formin-cappedfilaments, the mechanism remains to be determined. One pos-sibility is that profilin presents monomers in an orientationoptimal for their addition to barbed ends. It is thought thatonly about 2% of random collisions between an actin monomerand the free barbed end of a filament result in effective dockingand addition (83). FH1–profilin–G-actin interactions may in-crease the efficiency of this step. A second possibility, notmutually exclusive from the first, is that profilin-FH1 interac-tions may increase the concentration of G-actin near thebarbed end of the filament to increase the rate of elongation.The stage is set for testing these models. Another criticalaspect of the mechanism that should be tested is whether theFH1 directly transfers profilin-bound monomers to the barbedend, as suggested by a recent kinetic modeling study (380).

Bud6: stimulation of Bni1-mediated actin assembly. Bud6(also called Aip3) was first identified in both a yeast two-hybridscreen for actin-interacting proteins (13) and a genetic screenfor mutants defective in the bipolar budding pattern (426).Bud6 localization in cells is similar to that of Bni1 (14, 284),and bud6� cells show defects in formin-mediated actin cableassembly (99, 325). Further, in two-hybrid assays Bud6 hasbeen shown to interact with Bni1 (97) and Bnr1 (182). Subse-

quent biochemical work demonstrated that the C-terminal halfof Bud6 (i) binds specifically to actin monomers but not fila-ments, (ii) stimulates nucleotide exchange on G-actin, and (iii)enhances Bni1-mediated actin assembly in vitro additively withprofilin (253). Consistent with these effects, bud6� was foundto be synthetic lethal with pfy1-4, suggesting that Bud6 andprofilin have related and/or complementary cellular functions.Subsequently, the Bud6-binding site was mapped to a shortsequence in the C terminus of Bni1, overlapping with its DAD(251), raising the possibility that Bud6 may participate in ac-tivating Bni1 from an inhibited state (see below). AlthoughBud6 interactions with both Bni1 and Bnr1 were detected inthe two-hybrid studies above, biochemical assays could detectinteractions of Bud6 specifically with Bni1 and not Bnr1. Fur-ther, mutational analysis revealed that key residues in Bni1required for Bud6 binding are not conserved in Bnr1. Actinassembly assays confirmed this differential interaction; Bud6stimulates Bni1 activity by �2- to 3-fold but has no effect onBnr1 activity. Further mechanistic studies will be required todetermine precisely how Bud6 enhances Bni1-mediated actinassembly, which in principle could arise from enhanced nucle-ation and/or elongation.

Multiple lines of genetic evidence indicate additional cellu-lar functions for Bud6 beyond Bni1 stimulation. For example,bud6� cells have more severe defects in actin organizationthan bni1� cells, suggesting that Bud6 may contribute (di-rectly or indirectly) to the regulation of Bnr1 and/or otheractin organizing factors. Further, Bud6 has roles in captur-ing astral microtubules to regulate mitotic spindle orienta-tion (160, 338, 339) and in organization of the endoplasmicreticulum at the bud neck to compartmentalize mother anddaughter cells (226).

Rho GTPases. The localized assembly of yeast actin cablesand cytokinetic rings by Bni1 and Bnr1 necessitates tight spa-tial and temporal control of their activities. Thus, one criticalquestion is how Bni1 and Bnr1 actin assembly activities areup-regulated and down-regulated. Work on mammalian form-ins mDia1 and mDia2 has shown that they are auto-inhibitedby interactions between their amino-terminal DID motif andtheir carboxyl-terminal DAD region (8, 392) (Fig. 11A), andthese interactions abolish actin nucleation activity of the FH2(212). Many formins also contain an amino-terminal GBD adja-cent to the DID (211, 322). Direct binding of Rho GTPases to theGBD is proposed to release formins from an auto-inhibitedstate (8, 392). However, inhibition of mDia1 FH2 activity byDID is only minimally relieved by addition of RhoA (211, 212),suggesting other and/or more mechanisms may be required forformin activation.

The overexpression of N- and C-terminally truncated Bni1and Bnr1 constructs produces excess actin assembly in vivo (97,99, 304, 325), which has led to the suggestion that Bni1 andBnr1 may be auto-inhibited like mDia1 and mDia2. This isconsistent with conservation in Bni1 (and to a lesser degreeBnr1) of the key functional residues in DAD (8). However, itshould be noted that the critical DAD-binding residues in DIDare not conserved in Bni1 (203, 267) and that direct interac-tions between N and C termini have not been reported foreither Bni1 or Bnr1. This leaves open the question of whetherBni1 and/or Bnr1 is auto-inhibited, which should be addressed



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through biochemical analyses of full-length proteins.Regardless of whether Bni1 and Bnr1 are down-regulated by

auto-inhibition or alternative mechanisms (e.g., transinhibitionsimilar to that of Las17), Rho GTPases play key roles in yeastformin regulation in vivo. Six different Rho family GTPases areexpressed in budding yeast: Rho1, Rho2, Rho3, Rho4, Rho5,and Cdc42. Bni1 interacts with Cdc42, Rho1, and possiblyRho3 and Rho4 (97, 190), while Bnr1 binds Rho4 (164). Bni1localization depends on Cdc42 (284, 304). Further, Dong andcoworkers have addressed the functional importance offormin-Rho interactions by testing the ability of N-terminallytruncated (activated) Bni1 and Bnr1 constructs to suppressdefects caused by rho mutants at different stages of the cellcycle (79). Their work implicates Cdc42 in the control offormin-mediated actin assembly at bud emergence, Rho3 andRho4 during bud growth, and Rho1 during the response to cellstress. Another study implicates Rho1 in regulating Bni1-me-diated actin assembly at the cytokinetic ring (369). These stud-ies have made an important contribution by providing workingfunctional pathways for formin regulation by Rho GTPases,but they leave open the possibility that some of the observedeffects are not direct. Thus, biochemical evidence for the ef-fects of Rho GTPases on formin activity is needed to clarify thefunctional roles of these interactions.

Other Bni1 and Bnr1 ligands. A number of other cellularfactors interact with Bni1 and/or Bnr1, for which the functionalimplications of the interactions are poorly understood. Tef1(eEF1� homologue) is a highly abundant translation elonga-tion factor that interacts with Bni1 FH2 (residues 1328 to 1513)(374), binds to and bundles F-actin in vitro (76, 262), andcontributes to actin cable organization (132). Binding of a Bni1fragment to Tef1 disrupts Tef1 bundling of F-actin (374), butthe effects of Tef1 on Bni1 activity have not been investigated.Interestingly, overexpression of TEF1 induces formation ofsupernumerary actin cables and enlarged depolarized cells(261), similar to the effects of overexpressing dominant forminconstructs. As Tef1 is one of the most abundant proteins inyeast cells (55) and may link functions of the actin cytoskeletonand translational control, it will be of great interest to definethe effects of the Bni1-Tef1 interaction.

Additional factors that may affect formin function includeHof1, Spa2, and Smy1. Hof1, an SH3 domain-containing pro-tein of the PCH family (216), interacts with Bnr1 FH1 by thetwo-hybrid assay and localizes to the bud neck similarly to Bnr1(176). Genetic data also suggest that Hof1 may regulate Bnr1function at the bud neck (215, 378). Hof1 is proposed to me-diate septum formation to facilitate cytokinesis, rather thanactomyosin ring formation (33), and thus Hof1 could regulateBnr1-mediated cable assembly to promote septation. Anotherformin ligand, Spa2, appears to interact with Bni1, as Bnr1lacks the Spa2-binding domain (113) (Fig. 11A). Spa2 is re-quired for proper localization of Bni1 (284), but its potentialeffects on Bni1 activity have not been tested. The kinesin-related protein Smy1 interacts by two-hybrid assay with Bnr1FH2 (182), but no functional role for this interaction has yetbeen reported. Intriguingly, Smy1 also interacts with type Vmyosin Myo2 (28), providing a possible link between cableassembly by Bnr1 and transport on cables.

Organization and Stabilization of Actin Filaments

Bundling proteins. The distinct functions of cellular actinarrays require organization of actin filaments into specializedarchitectures (e.g., patches, cables, and rings). This is regulatedby proteins that bundle, cross-link, and/or stabilize filamentsinto specialized configurations. Below, we discuss the knownmechanisms and cellular functions of yeast actin-binding pro-teins that bundle and/or stabilize filaments: Sac6, Scp1, Iqg1,Crn1, Abp140, Tef1, Tef2, Tpm1/Tpm2, and Cap1/Cap2.

(i) Sac6. Sac6 has two actin-binding domains (ABDs), eachcomprised of a pair of calponin homology (CH) domains (Fig.12). True to its initial codiscovery by genetic (4) and biochem-ical (84) approaches, a deeper understanding of the Sac6-actininteraction has developed through the combination of geneticand biochemical studies. Initially, SAC6 was identified as anextragenic suppressor of temperature-sensitive act1 alleles, inwhich point mutations in Sac6 suppressed the growth defectscaused by point mutations in actin (4, 5). Independently, Sac6was isolated on F-actin affinity columns and shown to localizein vivo to patches and cables (84). Then, it was demonstratedthat Sac6 bundles F-actin in vitro and is required for properactin organization in vivo (3). Subsequent genetic and bio-chemical analyses demonstrated that Sac6 contacts residues onthe surface of actin subdomains 1 and 2 exposed on the “edge”of the actin filament (46, 154). Further, some of the defectsobserved in sac6� cells (e.g., synthetic lethality with abp1� andsla2�) are mimicked by mutations at the Sac6 contacts in actinsubdomains 1 and 2 (153, 329). To date, the cognate actin-binding surfaces on Sac6 have not been identified, despite theavailability of Sac6/fimbrin crystal structures to guide muta-tional analyses (123, 189).

All actin-bundling proteins cross-link filaments by one oftwo mechanisms: (i) having multiple ABDs or (ii) oligomer-ization. Sac6 bundles filaments by having two separate ABDs.High-resolution EM studies on Sac6-decorated filaments agreewith previous genetic work defining the Sac6-actin interfaceand provide a model for the cross-link (135, 136, 386). Untilrecently, it was thought that Sac6 was not regulated and that itstwo actin-binding heads were available to interact with fila-ments. However, recent crystal structures of A. thaliana and S.pombe Sac6/fimbrin reveal a “closed” conformation, in whichthe two ABDs fold in on each other, preventing exposure ofresidues hypothesized to mediate actin binding (189). Thissuggests that an induced structural shift may be required foractin binding to occur. Consistent with this possibility, distinctconformations for Sac6 have been observed by EM and crys-tallographic studies, but how such rearrangements might beregulated is not yet clear. One possibility is that the EF-handmotif in Sac6/fimbrin (Fig. 12) allows regulation by calciumand/or other signaling molecules.

While key features of the Sac6-actin interaction now areunderstood, the cellular function(s) of Sac6 remains unclear.For instance, Sac6 function in endocytosis has not been re-solved. Actin patches in sac6� cells are impaired in movinginward from the cell cortex (174), suggesting that Sac6 maycontribute to actin-based force generation, but the mechanismof this contribution is unknown. Further, how does a bundlingprotein associate with a network of branched filaments such asan actin patch? Perhaps Sac6 bundles filaments at specific



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stages in patch maturation, either prior to or subsequent to thestage characterized by EM where bundles are not obvious (314,423). Additionally, Sac6 bundling may be used to link patchesand cables during fast patch movement on cables (159). An-other possibility that should be considered is that the abilitiesof Sac6 to bundle and stabilize filaments may be separate.Thus, Sac6 could function primarily to stabilize rather thanbundle filaments in patches. Consistent with this model, par-tially purified actin patches isolated from sac6� cells are lessstable than those isolated from wild-type cells (423). To resolvethese issues, mutants that specifically impair each of the ABDsof Sac6 should be generated, and actin patch ultrastructure insac6� mutants should be analyzed by EM to determinewhether there are visible defects in filament organization.

The precise function of Sac6 in actin cables is also unknown.sac6� cells show diminished cable staining and rounded cellmorphologies, indicative of polarity defects (3). Thus, Sac6may bundle and/or stabilize the actin filaments in cables. How-ever, this has not been tested directly, for instance, by measur-ing rates of cable turnover in sac6� cells. Finally, a functionalrole for Sac6 at the actomyosin ring has not been investigated.S. pombe fimbrin localizes to the medial ring and functionsduring cytokinesis (412), raising the possibility that Sac6 func-tions similarly.

(ii) Scp1. Scp1 (a calponin/transgelin homologue) is the onlyother component of patches or cables (other than Sac6) thatcontains a CH domain; in addition, the bud neck componentIqg1 contains a CH domain (see below). Like Sac6, Scp1 lo-calizes to actin patches and bundles and stabilizes actin fila-ments in vitro (129, 405). Deletion of SCP1 exacerbates defectscaused by deletion of SAC6, further suggesting their functionalrelationship. However, biochemical dissection of Scp1 activityreveals clear mechanistic differences between these two bun-dlers. First, Sac6 bundles filaments through the use of twoABDs, each comprised of a pair of CH domains. However,Scp1 bundles filaments through C-terminal sequences distinct

from its CH domain (Fig. 12). This relatively small C-terminalregion of Scp1 is both required and sufficient for bundling butdoes not appear to oligomerize, leaving its bundling mecha-nism uncertain. One possibility is that this fragment harborstwo separate ABDs. Alternatively, actin binding may alter theconformation of Scp1 to promote oligomerization (similar toIQGAP [see below]). Scp1-induced actin filament bundles alsoare qualitatively distinct from Sac6-induced bundles, consistentwith their apparently divergent actin interactions.

Scp1 has not been visualized on actin cables or rings, but itsrelatively low abundance compared to Sac6 may preclude vi-sualization on those structures. Indeed, even the more abun-dant Sac6 shows only faint localization on cables (84). Thus, itis an open possibility that Scp1 and Sac6 function together toregulate cables. Scp1 also is linked to formation of reactiveoxygen species regulating yeast cell death (130), but it is not yetclear whether this function requires Scp1 interactions withactin. Key to understanding Scp1 cellular function will be de-termining the respective roles of its N-terminal CH domainand C-terminal actin-bundling domain. It is possible that theCH domain functions as a “locator” domain to facilitate activ-ity of the C terminus. Alternatively, the CH domain may me-diate interactions with other cellular factors. Thus, future workon Scp1 should (i) define which domains of Scp1 are requiredfor its localization and (ii) identify new Scp1 ligands.

(iii) Iqg1/Cyk1. A third CH domain-containing protein inyeast is Iqg1 (also called Cyk1), an IQGAP homologue. MostIQGAP family members have a CH domain at their N termi-nus, calmodulin-binding IQ motifs, and a C-terminal Ras-GAPdomain (RGD). The RGD mediates interactions with Cdc42but lacks the GAP activity originally proposed (Fig. 12) (228).Cdc42 and Iqg1 two-hybrid interactions have been reported(279), but interactions could not be detected biochemicallybetween Iqg1 and any of the yeast Rho GTPases (96, 341),leaving open whether Iqg1 is regulated in this manner. TheIQG1 gene is essential for yeast cell growth, and Iqg1 localizes

FIG. 12. Actin filament-bundling proteins. Schematics of each protein are drawn to scale. Abbreviations: CC, coiled-coil; CLR, calponin-likerepeat; COOH, carboxyl-terminal domain. Known actin-binding domains are underlined. Yeast IQGAP (Iqg1) is included because mammalianIQGAP bundles actin filaments; however, this activity has not yet been demonstrated for yeast Iqg1. It is unknown how Abp140 bundles actinfilaments, since its actin-binding domain(s) has not been defined.



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to the bud neck in a manner dependent on its IQ motifs andtheir interactions with Mlc1 (96, 217, 342). The terminal phe-notype of iqg1� cells suggests that it plays a critical role incytokinesis.

Iqg1 binds to F-actin through its N terminus, which containsthe CH domain (96, 341). Filament bundling by Iqg1 has notbeen tested. However, mammalian IQGAPs bundle filaments,and this activity is regulated by their interactions with Cdc42and calmodulin. Cdc42-GTP enhances bundling, possibly bypromoting IQGAP oligomerization (114), whereas calmodulininhibits bundling (26). One implication from these studies isthat IQGAP likely bundles via oligomerization, rather thanutilizing multiple ABDs on a single polypeptide chain. SinceIqg1 interacts with calmodulin and Mlc1 (341, 342), and evenpossibly Cdc42, it may be regulated in a similar manner, butthis awaits future tests.

What specific cellular function does Iqg1 perform to pro-mote cytokinesis? Truncation analyses show that the CH do-main contributes to assembly of the actomyosin ring, that theIQ motifs are required for Iqg1 recruitment to the bud neck(via interactions with Mlc1), and that the RGD facilitates con-traction of the actomyosin ring (341). This puts Iqg1 at thecenter of a fascinating network of interactions driving celldivision. However, the mechanisms by which Iqg1 facilitatesactin ring formation and subsequent ring contraction remainunclear. Does the CH domain interact directly with actin fila-ments to promote ring formation, or are there other ligandsinvolved? What molecular interaction(s) of the Rho-bindingdomain promotes ring contraction?

(iv) Crn1. Here, we discuss the actin-binding and -bundlingactivities of Crn1 (yeast coronin); its role in regulating theArp2/3 complex is described above. The N terminus of Crn1forms a �-propeller domain that mediates a high-affinity inter-action with F-actin; the C terminus contains a coiled-coil do-main (Fig. 12). Purified Crn1 oligomerizes via its coiled-coildomain and cross-links filaments into either loose arrays ofoverlapping short filaments or smooth bundles of longer fila-ments (128). Overexpression of Crn1 in yeast causes arrest ofcell growth, which can be suppressed by truncating either theABD or the coiled-coil domain. Crn1 localization to actinpatches also depends on both domains (161). Thus, Crn1 lo-calization and function may depend on its ability to bundleF-actin. Alternatively, this may reflect a requirement for Crn1interactions with the Arp2/3 complex, also mediated by thecoiled-coil domain (161). Since there appear to be separatepools of Crn1 in cells—about 50% of which is associated withthe Arp2/3 complex (161)—Crn1 may be performing multiplefunctions that involve its coiled-coil domain.

Deletion of CRN1 causes no obvious defects in cell growth,morphology, or actin organization, suggesting its functions areredundant with other cellular factors. However, Crn1 shows nogenetic interactions with other bundling proteins (Sac6, Scp1,and Abp140). Instead, crn1� displays synthetic defects in cellgrowth and actin organization with act1-159 and cof1-22. Thisraises the intriguing possibility that Crn1 may promote actinturnover rather than stabilization (128). These and other ques-tions regarding Crn1 cellular function need to be resolved.

(v) Abp140. Abp140 is one of only two actin-binding proteinsidentified in yeast with no obvious mammalian counterpart, theother being Bud6 (Table 1). Abp140 was first purified from

crude yeast cell extracts as a 140-kDa protein with actin-bun-dling activity (17). However, the ABP140 open reading framepredicts a protein with a calculated mass of 71 kDa. Thissuggests either that Abp140 has abnormal properties thatcause retarded migration on sodium dodecyl sulfate-polyacryl-amide gel electrophoresis gels or that it oligomerizes in amanner resistant to the denaturing conditions of sodiumdodecyl sulfate-polyacrylamide gel electrophoresis gels. Themigration of Abp140 on gel filtration columns is consistentwith oligomerization (17). Thus, while the ABDs of Abp140have not yet been identified, this protein appears to multim-erize, which offers a potential mechanism for bundling.Abp140 localizes to both actin patches and cables. However,abp140� cells display no defects in cell growth, morphology, oractin organization. Genetic interactions of Abp140 with otheractin-binding factors have not been tested, leaving its cellularfunction unclear.

Since its initial characterization, Abp140 has been used as aGFP-fused marker for studies monitoring actin cable dynamics(104, 159, 419). As multiple bundling proteins colocalize withpatches and cables (e.g., Abp140 and Sac6 and possibly Scp1),an important mechanistic question is whether these factorscompete for and/or cooperate in binding F-actin and helpconnect patches and cables. In addition, cable architecturelikely is shaped by the activities of multiple bundling factorsworking in concert.

(vi) Tef1 and Tef2. Tef1 and Tef2 are the yeast homologuesof eukaryotic translation elongation factor 1� (EF1�), andthey bear 81% sequence identity to their human counterpart(374). Tef1 and Tef2 are identical proteins, encoded by genet-ically redundant genes, that together are essential for cell via-bility (333). In addition to their well-documented roles in pro-tein translation, all EF1� family members examined (fromyeast to humans) bundle F-actin in vitro (67). In addition,Xenopus laevis and human EF1� proteins sever microtubules(347). Thus, Tef1 has the potential to coordinate protein trans-lation with functions of both the actin and microtubulecytoskeletons.

Filament bundling was first demonstrated for Dictyosteliumdiscoideum EF1� (418) and more recently for Tef1 (261). Thebundling mechanism of Tef1 likely involves two separateABDs, which have been mapped to two distinct 50-residuestretches, located at the N and C termini of Dictyostelium EF1�(218). EF1� bundles actin filaments into a unique architecturethat excludes other cross-linking proteins from binding (283).Point mutations that disrupt the ability of Tef1 to bundlefilaments, but leave its translation function unaffected, causedefects in actin cable organization in vivo (132). However, itremains unknown whether Tef1 localizes to cables or otherF-actin structures, and thus it is unclear how it contributes tocable organization. As discussed above, Tef1 interacts with theactin-nucleating FH2 domain of Bni1, and this interaction dis-rupts F-actin bundling by Tef1 (374). However, it remains to bedetermined whether this interaction affects the actin cableassembly-promoting activity of Bni1. Another reported ligandof Tef1 is the actin monomer-binding protein Srv2/CAP (seebelow) (417), but the in vitro and in vivo consequences of thisinteraction remain unknown.

Finally, it will be interesting to learn how the functions ofTef1 in regulating the actin cytoskeleton are coordinated with



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its classical role in translation. The role of EF1� in translationrequires GTP binding and hydrolysis (168, 312). F-actin bind-ing inhibits the abilities of EF1� to bind GTP but promotesGTP hydrolysis (90). Thus, actin binding may drive EF1� intothe GDP-bound form, inactive for translation.

Nonbundling Actin Stabilizers

(i) Tropomyosins Tpm1 and Tpm2. Budding yeast expresstwo tropomyosin isoforms, TPM1 and TPM2, which functionprimarily to stabilize the filaments in actin cables and theactomyosin ring. Intriguingly, they are the only proteins knownto localize exclusively to cables, i.e., not also label patches.Tpm1 and Tpm2 have genetically overlapping yet possibly dis-tinct cellular functions. TPM1 expression levels are �6 timeshigher than TPM2 levels in exponentially growing cells (81),which correlates with the relative severity of their phenotypes.Whereas tpm1� cells lack readily detectable cables, have cellmorphology defects, and grow slowly (219), tpm2� cells showno obvious defects in growth, morphology, or actin organiza-tion (81). Further, overexpression of TPM2 rescues most of thedefects caused by tpm1� (D. Pruyne and A. Bretscher, per-sonal communication). The observation that tpm1� and tpm2�mutations are synthetic lethal is consistent with genetic redun-dancy in their functions but also leaves open the possibility thatthey perform some specific roles, as suggested by their bio-chemical differences (below).

Tpm1 and Tpm2 display many of the defining biochemicalproperties of tropomyosin family members, including dimer-ization, divalent cation-dependent F-actin binding, and resis-tance to denaturation at high temperatures (90°C) (61, 81,220). However, Tpm1 and Tpm2 have different actin-bindingstoichiometries (1 Tpm1 dimer per 4.6 actin subunits; 1 Tpm2dimer per 3.6 subunits). Further, Tpm2 can displace Tpm1from actin filaments, but Tpm1 fails to displace Tpm2 fromfilaments. Additionally, low levels of Tpm1 may actually en-hance Tpm2 binding (81). This suggests that Tpm2 may havestronger affinity for F-actin and that its actin-binding interac-tion is qualitatively distinct from that of Tpm1. A future chal-lenge is to determine how these biochemical differences trans-late into different cellular functions. This will likely requirecareful analysis of cable dynamics in tpm1 and tpm2 mutantsand identification of differential genetic interactions for thesemutants.

As discussed earlier, the generation of a fast-acting temper-ature-sensitive strain, tpm1-2 tpm2�, greatly facilitated the dis-covery that tropomyosin is required for cable stability (306).Tpm1 and Tpm2 may stabilize cables by prohibiting depoly-merization/turnover factors, such as cofilin, from associatingwith filaments. Consistent with this view, filament binding bytropomyosin and cofilin are mutually exclusive in vitro and inCaenor habditis elegans muscle cells (30, 278). This is alsoconsistent with the observation that yeast patches and cablesare decorated differentially by cofilin (patches) and tropomy-osin (cables). As actin cables are highly dynamic structures thatundergo rapid filament turnover (19, 419), it will be importantto determine whether tropomyosin-actin interactions are reg-ulated. It seems likely that maintaining rapid turnover of cableswould require active mechanisms to displace these filamentstabilizers. Indeed, such mechanisms have begun to emerge;

Tpm1 (and likely Tpm2) is acetylated directly by the Mdm20-containing NatB acetyltransferase complex, and this modifica-tion promotes Tpm1 binding to F-actin and cable stability invivo (146, 292, 350, 351). This and other modifications, as wellas competitive inhibitor mechanisms, may be required to main-tain the delicate balance between cable stability and rapidturnover.

(ii) Capping protein. All of the filament-stabilizing factorsdiscussed above (e.g., Sac6, Scp1, and Tpm1/Tpm2) decoratesides of filaments and reduce rates of subunit dissociation asfilaments shorten from their barbed and/or pointed ends. How-ever, factors that directly target filament ends can also havetremendous stabilizing effects. It is difficult to find a moreclassic example of this than the CapZ/conventional cappingprotein family, of which yeast capping protein (Cap1/Cap2) isa member. These heterodimeric proteins tightly associate withand cap the barbed ends of actin filaments (396). Cap1/Cap2localizes to actin patches, and strains carrying deletions in theCAP1 and/or CAP2 genes show depolarized patches and di-minished cable staining (10, 11). The potential role of cappingprotein in regulating cable stability has not yet been addressed.However, recent studies show that capping protein plays animportant role in driving actin polymerization to facilitate slowactin patch movements (Fig. 4) (174, 185). This effect may beexplained by capping protein limiting the local concentrationof free barbed ends of filaments to “funnel” available actinmonomers to these barbed ends (i.e., those that are notcapped) (287). This increases the rate of actin polymerizationat these free ends, because the monomer pool is not depletedas rapidly and polymerization rates depend on monomerconcentration.

Another important function that capping protein provides inactin networks is regulation of average filament length. Byrapidly associating with free barbed ends of filaments, cappingprotein limits filament growth and thereby maintains a shortaverage filament length. This is a critical property for theorganization of dense actin networks comprised of many in-terconnected very short filaments, such as those found at theleading edge of motile cells, in the actin “comet tails” trailingintracellular vesicles and pathogens, or in yeast actin patches.Since capping protein is ubiquitously expressed and relativelyabundant, the assembly of alternative actin arrays comprised ofsomewhat longer filaments (e.g., filopodia, stress fibers, andcables) requires that the filament length restrictions imposedby capping protein be actively bypassed. This is achieved bytwo mechanisms. First, capping protein can be regulated at thecell cortex by direct interactions with PIP2, which disruptscapping protein associations with barbed ends (332). Second,factors such as formins or Ena/VASP associate with the grow-ing barbed ends of filaments (processive capping) and protectthose growing ends from capping protein.

What is the mechanism for capping filament barbed ends?The recently solved crystal structure of vertebrate CapZ re-veals that the two subunits, which have almost no similarity inprimary sequence, fold into remarkably similar structures thatintertwine (415). Each subunit has at its C terminus a short“tentacle” that protrudes from the main structural body of theprotein, and mutational analysis shows that both tentacles con-tribute to capping in vitro. Further, mutations in the Cap1tentacle cause cellular defects similar to those caused by the



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null mutations (186). Intriguingly, the Cap1 tentacle makes amuch stronger contribution to capping than the Cap2 tentacle(186, 397). Further, the Cap1 tentacle is sufficient to cap fila-ments, albeit with far lower affinity than intact capping protein(186). Ultimately, a full biochemical understanding of the cap-ping mechanism will require identification of the tentacle bind-ing sites on actin, which have only been modeled to date andnot determined experimentally (396).

Rapid Turnover of Actin Structures

Importance of actin turnover in vivo. Although much re-search attention has been placed on defining the cellular mech-anisms for assembling actin filaments within networks, therapid disassembly of filaments is equally important. Only bymaintaining actin networks in a state of dynamic flux, withindividual filaments undergoing regulated turnover, are cellsable to remodel their networks swiftly in response to internaland external cues (e.g., during cell motility and cytokinesis)and coordinate time-sensitive steps in complex actin-depen-dent processes (e.g., endocytosis). In addition, new growth ofactin filaments relies on the rapid and steady turnover of olderfilaments, which is required to replenish the monomer pool.Thus, reduced rates of filament turnover deplete the monomerpool and limit new growth.

All three major actin structures in yeast (patches, cables, andrings) undergo rapid turnover, as demonstrated by the loss ofall visible F-actin staining within 1 minute after treatment ofcells with Lat-A (19, 369). As Lat-A is reported to block newfilament growth but not actively disassemble filaments, thissuggests that the filaments in patches, cables, and rings un-dergo rapid turnover. The importance of rapid actin turnoverfor cell function was first demonstrated in studies employingthe cof1-22 conditional mutant (204). This mutation impairsthe actin disassembly activity of Cof1 in vitro and dramaticallyreduces in vivo turnover rates of actin filaments in patches.This study showed a correlation between reduced rates of actinturnover and defects in endocytosis and cell growth. Similarresults were reached in studies on act1-159, demonstrating acorrelation between reduced rates of actin filament disassem-bly in vitro and reduced rates of actin turnover in vivo, accom-panied by defects in endocytosis (27). Together, these studiesstrongly indicate that rapid turnover of actin is required forendocytosis and cell viability.

Most growth of actin networks in vivo occurs at the barbedends of filaments, and disassembly is thought to occur primar-ily by dissociation of subunits from the pointed ends of fila-ments. However, the rate of dissociation of subunits frompointed ends is very low (0.3 to 0.8 subunits per s). Since ratesof filament turnover in vivo are up to 100 times higher (428),this means that cells require active mechanisms for disassem-bling filaments. In yeast, as in other organisms, this is achievedthrough the coordinated activities of multiple actin-bindingproteins working in concert. Below we describe the mecha-nisms and functions of each yeast actin-binding protein knownto promote actin turnover. All of these proteins are highlyconserved and, therefore, their mechanisms are likely to beconserved.

Cofilin/ADF. Budding yeast has a single essential cofilingene (COF1), which encodes a small (16-kDa) protein with

properties similar to other members of the ADF/cofilin family(25, 162, 246). The primary cellular function of cofilin is toaccelerate actin filament depolymerization and turnover (204);however, the biochemical mechanism underlying this activity isnot yet fully understood. Cofilin binds to actin filaments andmonomers in vitro, in both cases preferentially interacting withADP-bound versus ATP-bound actin (141). Cofilin inducesfilament fragmentation, which structural studies suggest isachieved by twisting of the filament to weaken lateral contactsbetween actin subunits (39, 241). As cofilin binds cooperativelyto F-actin (140, 141), it may cluster at multiple sites along afilament, with nonuniform decoration leading to severingevents. The other reported activity of cofilin, acceleration ofsubunit dissociation from filament pointed ends (51), could inprinciple also stem from filament twisting. Importantly, sever-ing by cofilin has the potential to promote either assembly ordisassembly of filaments depending on the availability of actinmonomers in cells and how efficiently the new barbed endsgenerated by severing are capped.

What cellular function(s) requires the essential activities ofyeast Cof1? Cof1 localizes to actin patches and has been shownto be required for endocytosis (204, 205, 246). Although cof1mutants have not yet been studied using time-lapse imaging fordefects in patch maturation, it seems likely that cof1-22 mu-tants will show defects in early slow patch movements, similarto capping protein mutants. Requirements at this phase ofpatch development seem to mirror those of other systems thatdepend on rapid actin polymerization and turnover, such asListeria motility (221). In addition to its role in regulating actinturnover at patches, Cof1 may also play an unexpected role indriving actin cable dynamics. While Cof1 is detected specifi-cally on actin patches in wild-type cells (246), it localizes toboth patches and cables in aip1� cells (317). Further, recentfunctional studies show that cofilin (and Aip1) promotes therapid turnover of actin cables and that aip1� mutations sup-press the actin cable defects caused by deletion of TPM1 (276).Finally, a recent study showed that S. pombe Adf1 localizes toand functions in formation and maintenance of the actomyosinring (264). Thus, Cof1 has the potential to regulate turnover ofall the major actin structures in yeast cells: patches, cables, andrings.

One major open question about Cof1 is whether its activitiesare regulated in vivo. The ability of animal cofilins to interactwith actin is regulated tightly by phosphorylation of a con-served serine residue in the N terminus, subject to control by acascade of kinases and phosphatases (404). In yeast Cof1, anS4E (cof1-3) mutation of the analogous serine residue is lethal,suggesting that if phosphorylation occurred it would block ac-tin binding and cofilin function, similar to what is seen inanimal cells. However, no phosphorylation of endogenousCof1 (isolated from exponentially growing cells) could be de-tected, which has led to the suggestion that Cof1 may not beregulated (205). One intriguing possibility, however, is thatCof1 may be down-regulated specifically when cells are notactively growing, such as in stationary phase. Another potentialform of Cof1 regulation is pH change in the cytosol, as pHregulates the in vitro activities of cofilin/ADF proteins, includ-ing Cof1 (140, 141, 162), inhibiting filament disassembly activ-ity at pH levels of �7.5. However, it remains uncertain whetherpH is regulated locally in yeast cells and, therefore, whether



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this regulatory mechanism contributes to Cof1 function in vivo.Finally, the activity of ADF/cofilins is down-regulated by directinteractions with PIP2 (250, 272), which may have importantimplications for regulation of Cof1 at cortical sites, such asactin patches.

Aip1. Aip1 is an important binding partner and cofactor ofcofilin that promotes actin filament disassembly and turnover.Aip1 was first identified in a yeast two-hybrid screen for actin-interacting proteins (13) and later found as a cofilin-interactingfactor through biochemical affinity (275), two-hybrid (317),and genetic (163) approaches. Together, Aip1 and cofilin pro-mote net actin filament disassembly in vitro, and aip1� andcof1 mutations show genetic interactions (7, 244, 275, 317).Like cofilin, Aip1 localizes to actin patches (317), where it hasa demonstrated role in promoting rapid actin turnover (276).

Elucidation of the Aip1 mechanism has seen much recentprogress. Aip1 caps the barbed ends of cofilin-severed fila-ments to prevent filament reannealing and block elongation(23, 274) and enhances the filament-severing activity of cofilin(277). Structurally, Aip1 is comprised of two seven-bladed�-propellers linked by a short hinge domain (Fig. 13A) (245,384). Mutagenesis studies have defined two actin-binding sur-faces on Aip1 that are critical for its functions in vitro and invivo, one located on each �-propeller (66, 243, 245, 276). Thecofilin-binding surface lies in the hinge region of Aip1 locatedbetween the two actin-binding sites (66). The C-terminal actin-binding site specifically recognizes cofilin-bound F-actin (276).Since cofilin alters the twist of F-actin (241) and Aip1 pro-motes filament disassembly specifically in the presence ofcofilin, this C-terminal actin-binding site may recognize a cofilin-induced conformational state of F-actin. These properties tai-lor Aip1 for capping cofilin-severed filaments and, indeed,mutation of the C-terminal actin-binding site abolishes Aip1’scofilin-dependent activity. Taking these observations together,a model emerges for the Aip1 mechanism and function. Cofilindecorates the sides of filaments and helps recruit Aip1, whichbinds directly to F-actin and cofilin. Together, cofilin and Aip1induce filament severing to produce short fragments of F-actin(less than 50 nm) capped at their barbed ends by Aip1. Thesefragments disassemble rapidly by the dissociation of subunitsfrom their pointed ends.

Until recently, Aip1 and cofilin were thought to promoteactin turnover specifically at patches, owing to their localiza-tion to patches. However, as discussed in above (see “Actincable dynamics and turnover”), cofilin and Aip1 have recentlybeen shown to promote rapid turnover of cables (276). Therapid nature of the cofilin-Aip1 mechanism of disassemblingfilaments may explain why in wild-type cells cofilin and Aip1are not detected on cables. Cables are faint to begin with, andif only a small percentage of filaments are decorated withcofilin, it would be difficult to detect. In contrast, in aip1�cells, severing by cofilin produces uncapped barbed endsthat undergo new polymerization, consistent with cablethickening and detection of cofilin on cables. This also ex-plains why thickened cables decorated with cofilin are ob-served in the cof1-19 strain, a mutant that disrupts barbedend capping by Aip1 (317).

Srv2/CAP. Srv2 was initially identified in yeast as a suppres-sor of the activated rasV19 allele (Srv2) and as an adenylylcyclase-associated protein (CAP) (103, 107). The yeast protein

FIG. 13. Regulation of actin filament turnover by a network ofinteracting proteins. (A) Schematics of proteins discussed in the actinturnover section of the text. Abbreviations: �-propeller, forms �-pro-peller tertiary structure composed of WD repeats; CC, coiled-coil;�-helix, forms tertiary structure composed entirely of �-helices thatresembles 14-3-3 domains (201, 238, 425); P, polyproline domain;WH2, WASp homology 2; �-sheet, forms tertiary structure composedentirely of �-sheets (78). (B) Model for the sequential events in actinturnover. Cofilin (orange) binds cooperatively to and severs older(ADP-bound) actin filaments. Aip1 (green) interacts with cofilin andF-actin to cap the barbed ends of severed filaments. Aip1-cappedfilaments disassemble rapidly from their pointed ends, generating apool of cofilin-bound ADP-actin monomers that cannot exchange ATPfor ADP. The dodecameric Srv2 complex (blue), tethered to F-actinvia the SH3 domain of Abp1 (red), rapidly displaces cofilin and bindstightly to ADP–G-actin. This facilitates rapid nucleotide exchange(ATP for ADP) on actin. Srv2 has 100-fold-lower affinity for ATP–G-actin than ADP–G-actin and thus hands off ATP–G-actin to profilin.Profilin interacts directly with Srv2 and may accelerate nucleotideexchange on actin during this process. Profilin-bound ATP-actinmonomers are available for rapid addition onto the barbed ends offilaments. (C) Relay of actin monomers from cofilin to Srv2 to profilin,facilitated by their specific affinities for ADP- versus ATP-actin. Theproposed sequence of G-actin handoffs is boxed with arrows. Kd valuesare taken from references 36, 51, 192, 237, 379, and 383.



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was named Srv2, but homologues in other species are calledCAP. Yeast Srv2 interacts with the adenylyl cyclase complexthrough a binding site in its N terminus, whereas the C termi-nus of the protein binds to actin monomers and plays a centralrole in regulation of actin dynamics in cells (Fig. 13A) (112,117, 180, 237, 424). The adenylyl cyclase-binding activity doesnot appear to be conserved in animals and plants, but its rolein regulating the actin cytoskeleton is conserved in all Srv2/CAPs tested so far (158). Deletion of the SRV2 gene in yeastcauses defects in cell growth and actin organization, and ex-pression of CAP homologues from diverse species rescuesthese defects. In addition, srv2� defects are suppressed par-tially by overexpression of profilin, suggesting a relationshipbetween their in vivo functions (385). Until recently, the spe-cific function performed by Srv2/CAP in the actin cytoskeletonwas unknown.

Recent studies have revealed that Srv2/CAP promotes therapid turnover of actin networks by recycling newly depolymer-ized actin monomers and cofilin (23, 31, 237, 249). After cofilinpromotes filament disassembly, it remains bound to ADP-actinmonomers and inhibits nucleotide exchange (ATP for ADP)(36, 205, 269). This leads to the accumulation of a pool ofcofilin-bound ADP-actin monomers, blocked from returning toan ATP-bound assembly-competent state. Therefore, cells re-quire a mechanism to displace cofilin from ADP-actin mono-mers. Classical models depict profilin performing this functionthrough competition (295). However, profilin fails to competeeffectively with cofilin for binding to ADP–G-actin, suggestingthat a “middle man” is required to facilitate exchange of actinmonomers from cofilin to profilin. Srv2/CAP has been demon-strated to perform this function in vitro (23). The C terminusof Srv2/CAP binds to and promotes nucleotide exchange onactin monomers. Further, full-length Srv2 catalyzes the con-version of cofilin-bound ADP-actin monomers to profilin-bound ATP-actin monomers. Thus, Srv2/CAP promotes actinturnover by (i) recycling cofilin for new rounds of filamentdisassembly and (ii) facilitating nucleotide exchange on G-actin to replenish the assembly-competent monomer pool (Fig.13B) (23, 249).

More-recent studies have begun to define the mechanismunderlying these effects (237). The C terminus of Srv2 has an�100-fold-higher affinity for ADP–G-actin (Kd 0.02 �M)than ATP–G-actin (Kd 1.9 �M), which enables it to competefor ADP–G-actin binding with cofilin (Kd 0.02 to 0.1 �Mcofilin) (36, 51, 237, 379). Further, the N terminus of Srv2 mayassist in these events, as it binds specifically to cofilin–G-actincomplexes and is required for rapid conversion of cofilin-bound ADP–G-actin to profilin-bound ATP–G-actin (23, 249).In this respect, it will be interesting to learn whether the N ter-minus of Srv2 has the ability to weaken cofilin-ADP-actin mono-mer interactions. Following transfer of ADP–G-actin from cofilinto Srv2, rapid nucleotide exchange occurs on actin. However,Srv2-bound ATP-actin monomers are not competent for addi-tion onto filament barbed ends (237). This means that anotherhandoff is required (to profilin). Addition of profilin to Srv2-bound G-actin releases monomers for addition onto barbedends, suggesting a transfer has occurred (237). Thus, Srv2/CAPcatalyzes conversion of actin monomers from the ADP- toATP-bound state and, like many enzymes, Srv2 has a signifi-cantly higher affinity for its substrate (ADP-actin) than its

product (ATP-actin), which accelerates the reaction.One of the most fascinating properties of Srv2/CAP, with

likely implications for its cellular function and mechanism, isthat it hexamerizes to form a high-molecular-weight complex.Native Srv2 complex isolated from yeast cells is �600 kDa,with a sedimentation coefficient of 15S and dimensions of �15by 30 nm long as determined by rotary shadowing EM ofpurified complexes (23). The complex is comprised entirely oftwo proteins, Srv2 and actin, present at a 1:1 molar stoichiom-etry, suggesting a heterododecamer (assembled from six Srv2and six actin molecules). Each Srv2 molecule in the complexcan interact with G-actin and at least three actin-binding pro-teins: cofilin, profilin, and Abp1. Binding of cofilin and profilinmay promote the handoffs described above. Binding of Abp1 isrequired for Srv2 localization to yeast actin patches (214) andtethers Srv2 complex to actin filaments in vitro (23). Since theAbp1-binding site on Srv2 is located in its C terminus, proximalto the G-actin-binding domain, it will be interesting to learnwhether Abp1 also affects Srv2 activity. Taken together, theseinteractions suggest that the Srv2 complex may serve as amolecular hub to facilitate dynamic interactions among multi-ple proteins in promoting actin turnover. This concept raisesimportant questions about the structure and physical proper-ties of the complex and how they influence function. For in-stance, what is the nucleotide-bound state of the actin subunitsbound to the complex? Do these subunits freely exchange withactin monomers in solution? How are Srv2 and actin moleculesarranged within the complex? Is multimerization of Srv2 re-quired for its activities and in vivo functions?

Profilin. Profilin is a small (�15-kDa), ubiquitous, abundantactin monomer-binding protein, the biochemical activities ofwhich have long been studied (409). In all organisms examined,mutation of profilin is lethal and/or causes severe defects inactin organization and cell morphology. In S. cerevisiae, a de-letion mutant (pfy1�) and point mutations that impair actinbinding (e.g., pfy1-4) show that profilin is critical for cellgrowth, morphology, and actin organization (134, 410).

Association of profilin with actin monomers (Fig. 8) has fourknown effects of potential importance in vivo. First, it sup-presses spontaneous actin assembly by sterically hindering in-teractions between actin monomers required for the formationof polymerization intermediates (295). Second, it restricts actinmonomer addition to filament barbed ends, blocking additionto pointed ends (297). Third, it can increase the rate of elon-gation at filament barbed ends capped by formins (195, 321).Fourth, it accelerates nucleotide exchange (ATP for ADP) onG-actin (242). In biomimetic assays for cell motility that em-ploy purified proteins, profilin becomes rate limiting for actinassembly, consistent with its proposed function in acceleratingactin turnover (206, 221, 230). However, it remains unclearwhich activities of profilin are required in vivo.

Synthetic lethal interactions between pfy1-4 and cof1-22,srv2�, and act1-159 are consistent with profilin promoting actinturnover in vivo. However, it is important to note that directevidence for this proposed function is still lacking, as actinturnover rates have not been measured for profilin mutants invivo. One hypothesis is that profilin’s ability to promote nucle-otide exchange on G-actin is critical for its proposed turnoverfunction. This is supported by the observation that cellulardefects caused by the pfy1-4 allele (impaired in binding actin)



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are suppressed by act1-157, an allele with an increased intrinsicrate of nucleotide exchange (410). This study concluded thatprofilin has a critical role in vivo in accelerating actin monomerrecycling. However, other observations suggest that profilinmay contribute to actin turnover by a mechanism other thanpromoting nucleotide exchange on G-actin. First, S. cerevisiaeprofilin increases nucleotide exchange on G-actin by only �3-fold in vitro, orders of magnitude less than human and birchpollen profilin (88). Second, there are no mutants yet availablethat impair nucleotide exchange without disrupting actin bind-ing. Thus, it is not possible to separate effects of profilin onnucleotide exchange from its other effects, such as promotionof actin assembly by delivering monomers to the ends of Ena/VASP- or formin-capped filaments. Third, Arabidopsis profilin,which binds to G-actin without promoting nucleotide exchange(290), complements pfy1� yeast cells (62); this raises the pos-sibility that acceleration of nucleotide exchange may not be acritical function of profilin in yeast.

What turnover function might profilin perform beyond ac-celerating nucleotide exchange on G-actin? Recent studies(see “Srv2/CAP” above) suggest that profilin plays a key rolealong with Srv2/CAP in facilitating the rapid conversion ofactin monomers (from a cofilin-bound ADP–G-actin state toan assembly-competent profilin-bound ATP–G-actin state). Inthis proposed mechanism, profilin binds to G-actin followingnucleotide exchange. This is consistent with (i) profilin havingvery low affinity for ADP–G-actin and significantly higher af-finity for ATP–G-actin (383) and (ii) profilin failing to competewith cofilin for ADP–G-actin binding (23). In cells, profilinmay first bind to ATP-actin monomers while they are stillassociated with Srv2/CAP, just after they have undergone nu-cleotide exchange (see above). Srv2/CAP has a weak affinityfor ATP–G-actin, which facilitates the handoff to profilin(237). In addition, since Srv2-bound actin monomers cannot beadded to filament barbed ends, an additional handoff is re-quired to factors such as profilin that readily promote barbedend assembly. This model is supported by the ability of profilinto siphon ATP-actin monomers from Srv2/CAP and makethem available for barbed end growth (237). It is also sup-ported by physical interactions between profilin and Srv2/CAPthat may facilitate monomer handoffs (32, 82). Thus, the loss ofprofilin would stall this mechanism and rapidly deplete theassembly-competent pool of actin monomers.

Another point to consider is that acceleration of nucleotideexchange by profilin requires its interaction with actin but notproline-rich sequences, yet mutations in profilin that impairpolyproline binding (without affecting actin binding) cause se-vere defects in cell growth and actin organization (134, 225,410). This suggests that profilin function in vivo depends oninteractions with both actin and polyproline targets, indicatingthat profilin must have critical functions beyond nucleotideexchange on actin. What is the other function of profilin invivo? Profilin targets actin monomers to the growing barbedends of filaments associated with Ena/VASp and formins. In S.cerevisiae, which does not express Ena/VASP homologues, pro-filin is believed to target actin monomers to barbed ends offilaments that the formins Bni1 and Bnr1 assemble into cablesand the actomyosin ring. Profilin binding to the proline-richformin FH1 domain is required for both Bni1 and Bnr1 to

assemble filaments from profilin-actin monomers in vitro (251,300, 326). Consistent with this function being important invivo, profilin mutants impaired in binding polyproline displaysevere defects in actin cable assembly and polarized cell growth(134, 225, 410).

Twinfilin. Twinfilin is a widely conserved 40-kDa proteincomprised of two tandem ADFH domains (Fig. 13A). Thefounding member of the twinfilin family (Twf1) was identifiedin S. cerevisiae by mining the genome database for cofilin ho-mologues and by isolation from crude extracts on F-actin af-finity columns with other actin-binding proteins, including cap-ping protein (125). Initial biochemical characterization of Twf1showed that it is an actin monomer-sequestering protein. Al-though twf1� mutants showed normal cell growth and mor-phology, their actin patches were slightly brighter, consistentwith actin turnover defects, and recent work has demonstratedthat loss of TWF1 reduces rates of actin patch turnover (252).twf1 mutants also are synthetic lethal with cof1 and pfy1 mu-tants (125), consistent with a role in promoting actin turnover.

Biochemical analyses have left the precise role of Twf1 inactin turnover somewhat enigmatic. Twinfilin sequesters actinmonomers (125), with a strong binding preference for ADP–G-actin (Kd � 0.05 �M) over ATP–G-actin (Kd � 0.5 �M)(273). While each ADFH domain of twinfilin is capable ofbinding G-actin, the C-terminal ADFH domain binds to G-actin with affinity similar to that of intact twinfilin, suggestingthat the N-terminal ADFH domain may have an alternativefunction. Twf1 also binds directly to capping protein (Cap1/Cap2) and requires capping protein for its localization to actinpatches in vivo and for genetic functions shared with cofilin(101, 285). However, the Twf1-capping protein interactiondoes not affect the capping activity of Cap1/Cap2 or the mono-mer-sequestering activity of Twf1. Thus, two discrepancies re-garding Twf1 function have been the following: how do Twf1-actin interactions promote actin turnover in vivo, and whatfunction is served by the Twf1-capping protein interaction?

Two recent studies have shed new light on these issues. Inone study, mammalian twinfilin was demonstrated to cap fila-ment barbed ends; however, this activity was not conserved inyeast Twf1 (143). In a second study, yeast Twf1 was found tobind to and sever actin filaments at low pH (252), an intriguingresult given the known pH dependence of ADF/cofilin activi-ties on F-actin (24, 85, 162). Further, capping protein directlyinhibits the filament-severing activity of Twf1 (252), represent-ing the first biochemical function ascribed to this physiologicalinteraction. Presently, it is difficult to explain the in vivo sig-nificance of the low-pH dependence of filament-severing ac-tivity in vitro. One possibility is that posttranslational modifi-cations of Twf1 in vivo enable it to sever filaments at moreneutral pH levels. Alternatively, Twf1-severing activity may beregulated by pH fluctuations in vivo.

In summary, twinfilin function on actin turnover is likely toinvolve a balance among its interactions with G-actin and F-actin, with capping protein regulating the balance betweenthese activities. Additionally, Twf1 binds to PI(4,5)P2, whichinhibits its interactions with both G-actin (285) and F-actin(252), suggesting that Twf1 function requires coordinated reg-ulation by multiple signals.



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CLOSING REMARKS AND FUTURE DIRECTIONS

Over the past decade, intense research efforts have shownthat the three most prominent actin structures (patches, cables,and rings) are all highly dynamic, with patches and cablesassembling and turning over in less than 60 seconds. Further,numerous genetic, two-hybrid, and biochemical studies havebrought us close to having a complete list of actin-associatedproteins involved in these processes. For many, their activitieson actin have been determined (Table 1) and their atomicstructures now are available (e.g., Abp1, the Arp2/3 complex,cofilin, coronin, profilin, Sac6, Srv2/CAP, and twinfilin). All ofthis sets the stage for a detailed mechanistic dissection of theelegantly choreographed events in actin network assembly andfunction.

One key future challenge is to develop new biochemicalstrategies for defining the activities of actin-associated proteinsin cellular mixtures. To date, the effects of actin-binding pro-teins on actin have been studied primarily using in vitro assayscontaining one or two purified actin-binding proteins and ac-tin, and often these components are not present at concentra-tions or molar ratios found in vivo. However, the activities ofactin-binding proteins can be affected strongly by their in vivobinding partners (e.g., Crn1-Arp2/3, Aip1-cofilin, and Srv2/CAP-cofilin). Most actin-binding proteins do not functionalone but rather in stable and/or dynamic complexes. Further,actin-binding proteins must compete for limited binding siteson actin, which can greatly influence their ability to affect actindynamics and organization. Thus, the activities of actin-bindingproteins should be examined in complex mixtures that moreclosely reflect physiological conditions. Some of these com-plexes can be isolated from yeast cells in native form; otherscan be “built up” from purified proteins with known interac-tions. This also emphasizes the need to define local cellularconcentrations and in vivo molar ratios of actin-binding pro-teins, as has been done recently for some actin-binding pro-teins in fission yeast (413).

A second future challenge is to understand how cellularsignals regulate the activities of actin-binding proteins toalter actin filament assembly, organization, and turnover.Live cell imaging studies have revealed that the assemblyand disassembly of yeast actin networks (patches, cables,and rings) are subject to exquisite spatial and temporalregulation. Further, many signaling molecules have beenimplicated in regulating actin cytoskeleton organization andfunction (e.g., Pkc1, Ark1/Prk1, Ste20/Cla4, Cdc28, and mi-togen-activated protein kinases). However, in the vast ma-jority of cases the actin-binding protein targets of thesesignaling pathways are unknown, and therefore the mecha-nisms of regulation are unclear. One of the few examples ofdirect regulation recently demonstrated is phospho-regula-tion of Pan1 NPF activity by Prk1 kinase (371). Additionally,Rho GTPases directly regulate actin-binding proteins suchas Bni1 and Bnr1 and control Ste20/Cla4 kinases and thepolarisome scaffold protein Bem1 (170). More studies ofthis nature are needed, as these examples likely representonly the tip of the iceberg for signaling molecules governingthe activities of actin-binding proteins.

ACKNOWLEDGMENTS

We are very grateful to all who contributed to the assembly andediting of this review. We are especially grateful to Jacqueline Hendriesfor her devoted assistance during the early stages and to the followingpeople for their generous help in editing the review prior to submis-sion: S. Buttery, F. Chang, K. Daugherty, M. Gandhi, K. Kono, S.Martin, K. Okada, D. Pruyne, A. Rodal, and I. Sagot. We also thankthe reviewers for valuable suggestions and D. Amberg, D. Botstein, A.Bretscher, M. Eck, J. Pringle, L. Pon, A. Rodal, and M. Rosen for theirpermission to reproduce original images. Finally, we apologize for theomission of important contributions made to this field that we mayhave overlooked in writing this review.

This work was supported by a grant from the NIH (GM63691) toB.L.G.

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