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Microbiol Mol Biol Rev. 2006 Sep; 70(3): 605–645.
PMCID: PMC1594590

The Yeast Actin Cytoskeleton: from Cellular Function to Biochemical Mechanism


All cells undergo rapid remodeling of their actin networks to regulate such critical processes as endocytosis, cytokinesis, cell polarity, and cell morphogenesis. These events are driven by the coordinated activities of a set of 20 to 30 highly conserved actin-associated proteins, in addition to many cell-specific actin-associated proteins and numerous upstream signaling molecules. The combined activities of these factors control with exquisite precision the spatial and temporal assembly of actin structures and ensure dynamic turnover of actin structures such that cells can rapidly alter their cytoskeletons in response to internal and external cues. One of the most exciting principles to emerge from the last decade of research on actin is that the assembly of architecturally diverse actin structures is governed by highly conserved machinery and mechanisms. With this realization, it has become apparent that pioneering efforts in budding yeast have contributed substantially to defining the universal mechanisms regulating actin dynamics in eukaryotes. In this review, we first describe the filamentous actin structures found in Saccharomyces cerevisiae (patches, cables, and rings) and their physiological functions, and then we discuss in detail the specific roles of actin-associated proteins and their biochemical mechanisms of action.


All cells undergo rapid remodeling of their actin networks to regulate such critical processes as endocytosis, cytokinesis, cell polarity, and cell morphogenesis. These events are driven by the coordinated activities of a set of 20 to 30 highly conserved actin-associated proteins, in addition to many cell-specific actin-associated proteins and numerous upstream signaling molecules. The combined activities of these factors control with exquisite precision the spatial and temporal assembly of actin structures and ensure dynamic turnover of actin structures such that cells can rapidly alter their cytoskeletons in response to internal and external cues. One of the most exciting principles to emerge from the last decade of research on actin is that the assembly of architecturally diverse actin structures is governed by highly conserved machinery and mechanisms. With this realization, it has become apparent that pioneering efforts in budding yeast have contributed substantially to defining the universal 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 then we discuss in detail the specific roles of actin-associated proteins and their biochemical mechanisms of action. For more detailed information on actin-based cellular processes, we refer readers to a number of excellent recent reviews that cover these topics in greater depth: endocytosis (93), mitochondrial inheritance (42), vacuolar inheritance (45), establishment of cell polarity (302, 305, 334), and cytokinesis (22, 33). In some sections we present data and concepts in a historical order, where tracing the progression is instructive. We also note that the majority of the cellular actin structures and their components are conserved between budding yeast and fission yeast, and in many cases insights from fission yeast preceded those from budding yeast. Due to space constraints we have focused the review on the budding yeast actin cytoskeleton, but we highlight experiments from fission yeast to reinforce specific points.


First Contact

The first major insights into organization of the yeast actin cytoskeleton 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 been established that actin was expressed from a single essential gene in S. cerevisiae, ACT1 (349), but the distribution of actin in yeast cells remained a mystery. Using fluorochrome-labeled drugs (2) and actin antibodies (183), Adams and coworkers visualized filamentous actin in chemically fixed cells and described an “unusual distribution of actin” consisting of two distinct structures (Fig. (Fig.1):1): cortical spots (“patches”) and long fibers (“cables”). Their elegant description of actin organization (see details below) set the stage for the recent advances in cell biological analyses on yeast actin described in the following sections: “Numerous small patches or spots were seen distributed over the cell surface; the cortical location of these spots was clear when the microscope was focused up and down. In addition, fibers were seen coursing through the cells, generally in a direction roughly parallel to the long axes of the cells. In unbudded cells, there was often a concentration of fluorescence near one pole, and in cells with very small buds there was generally a pronounced concentration of fluorescence in the bud. In both cases, the concentration of fluorescence was usually resolvable as a cluster of individual spots. In cells with medium-sized buds, there was typically a higher concentration of fluorescent spots in the bud than in the mother cell. Cells with very large buds often showed a concentration of fluorescence in the neck region, sometimes apparently as clusters of spots and sometimes apparently as a band.” (Reproduced with permission from reference 2.)

FIG. 1.
Cell cycle-regulated organization of the S. cerevisiae actin cytoskeleton. Yeast cells at different stages in the cell cycle contain three visible F-actin structures: cortical actin patches, polarized actin cables, and a cytokinetic actin ring. While ...

In addition to patches and cables, Adams and coworkers noted the presence of a third actin structure located at the neck of some large-budded cells, which they referred to (above) as a “band.” Twenty years later, it is clear that this was the first sighting of the budding yeast actomyosin contractile ring. However, the concept of a budding yeast actomyosin ring lay relatively dormant over the next 14 years, in part due to the fact that actin rings are present only transiently during the cell cycle and therefore observed in only a limited number of cells in an asynchronously growing culture and in part because they can be more difficult to image than patches and cables. In addition, many in the field believed that the narrowness of the bud neck, approximately 1 μm during budding yeast cytokinesis (327), precluded the requirement for an actomyosin contractile ring. Then, in 1998, the presence of a functional actomyosin ring was confirmed by two independent studies (34, 217) which revealed that the events of cytokinesis in yeast and animals were more similar than previously thought.

Studies in 1986 revealed that the three visible F-actin structures in budding yeast—patches, cables, and rings—are also present in the fission yeast Schizosaccharomyces pombe (231). A large body of literature since then has shown that the mechanisms regulating patch and cable assembly in fission and budding yeast are similar but that the rules governing assembly of the actomyosin ring differ significantly (see below). In addition, smaller and/or less visible actin structures may exist in both fission and budding yeast, as suggested by the requirement for actin dynamics in the processes of vacuolar fusion (92), endoplasmic reticulum cortical dynamics (301), and chromatin remodeling (38). Visualizing these actin structures in vivo represents a key challenge for the future.

Actin Patches and Endocytosis

Early observations.

Initial observations showing that patches are concentrated at sites of polarized growth suggested that patch function might be linked to secretion. However, patches are decorated with numerous actin-associated proteins, and as mutations in these patch components were characterized, it became apparent that their primary defects were endocytic rather than secretory. Multiple patch components (e.g., actin, Arc35, Rvs167, Sac6, Sla2, and Vrp1) were identified in random screens for mutants defective in endocytosis, end mutants (64, 86, 93, 202, 260, 309, 311). In addition, an endocytic function 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 to steadily draw correlations between the functions of patch components and endocytosis, but this linkage was not solidified and fully accepted until recent light and electron microscopy (EM) studies addressed the dynamic movement and ultrastructure of cortical actin patches (see below).

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

Actin patch dynamics.

An important and unanticipated property of actin patches was discovered with the advent of green fluorescent protein (GFP) fusions to monitor real-time behavior of proteins in cells. By tagging patch components (Sac6, Cap1/Cap2, Abp1, and actin) with GFP, two initial studies demonstrated that patches are highly motile (80, 387). These and other early studies showed that the lifetime of actin patches is approximately 10 to 20 seconds and that actin patches first assemble at sites of polarized cell growth and then move slowly and nondirectionally along the cell cortex. In addition, more-rapid movements were observed. Patch motility rates ranged from 0.1 to 0.5 μm/s (52, 354, 387).

These observations raised an important mechanistic question: what provides the force driving patch movement? A decade ago, when patch motility was first observed, most forms of actin-based motility were thought to be myosin dependent. So, it came as a surprise when it was reported that the rate of patch motility was unaffected by mutations in any of the five yeast myosin genes: MYO3 and MYO5 (type I), MYO1 (type II), or MYO2 and MYO4 (type V) (354, 387). Instead, studies using Lat-A suggested that the actin filaments in patches undergo rapid turnover (19) and that Lat-A inhibits patch movement (52, 289), suggesting that actin polymerization may provide the force required for patch motility. As such, parallels were drawn between yeast actin patch motility and the actin polymerization-based motility of the intracellular pathogen Listeria monocytogenes, 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 approximately 0.4 μm/s (70, 368). While it has been appealing in models to depict actin patches like miniature Listeria actin comet tails trailing endocytic vesicles, patches have at least two key properties that distinguish them from Listeria. First, cofilin activity, and therefore rapid actin turnover, is required for Listeria motility (287) but not rapid patch movement (204, 287). Second, Listeria motility is autonomous in living cells and cell extracts, requiring only the actin tail that it forms for propulsion. However, the rapid phase of patch motility (see below) relies on an additional separate network of filamentous actin structures, actin cables (159).

From a number of recent studies, it has become apparent that actin patches mature in stages corresponding to different stages of endocytosis and characterized by different types of movement. One of the first indications of this behavior came from a study examining the dual localization and fluorescence resonance energy transfer between CFP-Abp1 (an activator of the Arp2/3 complex) and YFP-Sla1 (an early endocytic protein) (391). This showed that at any given time, a subset of patches colabeled with Abp1 and Sla1, while others contained exclusively Sla1 or Abp1. Intriguingly, the patches labeled exclusively with Abp1 were highly motile, whereas those labeled exclusively with Sla1 exhibited little movement. The authors hypothesized that “Sla1-only” patches might contain endocytic machinery and the arrival of Abp1 (and possibly other factors) facilitated actin polymerization-based rapid movement of patches. This study emphasized the need to consider temporal changes in the development and lifetime of an actin patch.

Subsequent studies by Kaksonen et al. (173, 174) resolved many of the key events in patch development by elegantly correlating temporal changes in patch motility with the arrival of specific components. Using pairs of integrated functional fluorescent tags on different patch components (Las17, Sla1, Sla2, Pan1, Abp1, and the Arp2/3 complex) and computer algorithms to track patch movements, they defined three stages in the lifetime of a patch. First, nonmotile patches, which contain Las17, Sla1, and Pan1, but not actin, form at the cell cortex. Next, polymerized actin appears, which coincides with the onset of slow patch movements at the cortex (0.05 to 0.1 μm/s). Patches transition to a phase of rapid inward movement from the cell cortex, likely as vesicles coated with actin filaments. Addition of Lat-A abolishes both the slow and fast patch movements (stages 2 and 3), suggesting that this motility is actin polymerization based (289, 407). The roles of specific patch components are discussed below in a model for patch development.

Shortly after the initial Kaksonen study (173), Huckaba et al. (159) reported three key findings clarifying patch function and mechanism. First, they showed that endocytic vesicles (marked by the lipophilic marker FM4-64) colocalize with motile patches (marked by Abp1-GFP) during slow and fast phases of movement (Fig. (Fig.2A).2A). This provided direct evidence that actin patches are sites of endocytic uptake. Second, they showed that rapid actin patch movement is mediated by polarized actin cables, as internalized actin patches colocalize with cables (Fig. (Fig.2B)2B) and move directionally with cables at a similar rate (∼0.3 μm/s), and rapid patch movements are lost upon disruption of cables by specific conditional mutants (159). Third, internalized patches/vesicles appear to fuse eventually with endosomal compartments and shed their actin coats. The movement of patches on cables is consistent with an earlier live cell imaging study from fission yeast (289). Thus, slow patch movements in the cell cortex (possibly to facilitate vesicle formation and scission) depend on an Arp2/3 complex-based actin polymerization mechanism, whereas rapid inward movement of patches depends on a mechanism of transporting patches on cables. These findings raise further questions that remain unresolved. What factors link patches to cables? How do cables deliver patches to endosomes, and are endosomes linked to cables? What signal triggers actin coat shedding at endosomes?

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 ...

In animal cell endocytosis, short bursts of actin polymerization at the cell cortex (similar to actin patch assembly) are accompanied by clathrin coat formation. Several yeast actin patch components are physically linked to clathrin and its adaptors, suggesting that clathrin may participate in patch dynamics (21). However, until recently, the direct involvement of clathrin in yeast endocytosis remained controversial, in part because endocytic defects in clathrin mutants are not fully penetrant, and in part because clathrin localization at cortical patches had not been demonstrated (21). In addition, clathrin-coated pits had not been observed in EM studies on the yeast cell cortex (255, 314), but two recent studies have provided convincing evidence that clathrin indeed localizes to early patches to facilitate endocytosis (174, 266). The localization of clathrin at the cell cortex had been masked by abundant clathrin staining on internal membranous structures. This technical limitation was overcome using total internal reflection fluorescence (TIRF) microscopy, which reduces fluorescent background and permits focused observation of fluorescence at the cell cortex. Both studies used TIRF to show that clathrin is recruited to early patches, possibly by Sla2, and then later disappears upon transition of patches to fast inward movement (174, 266). Further, it was shown that almost all clathrin patches at the cortex subsequently recruit actin (i.e., become actin patches) and that clathrin mutants cause delays in early stages of patch development. These studies have ended the debate over whether clathrin functions in yeast endocytosis and raise exciting new questions, such as what signaling mechanisms control recruitment and subsequent loss of clathrin at patches. Another factor that plays a key role in endocytosis in animal cells is dynamin, a GTPase that tubulates membranes and facilitates vesicle scission (298). Although multiple dynamins are expressed in budding yeast, their involvement in endocytosis remains to be demonstrated (93).

Actin patch ultrastructure.

Ultimately, to fully understand the inner workings of actin patches, a detailed understanding of patch ultrastructure will be required at a resolution beyond the limits of light microscopy. However, only a few studies using EM have been successful in elucidating details of patch ultrastructure, due to the unique technical challenges presented when analyzing actin in yeast cells: the presence of a cell wall, a concentrated and thus crowded cytoplasm, and a low concentration of actin filaments. Initial EM observations of two-dimensional thin slices revealed that actin patches associate with finger-like invaginations (potential sites of endocytosis) at the plasma membrane (256) (Fig. 3A and B). Additional work on thin slices demonstrated limited colocalization of actin with sites of receptor-mediated endocytosis (255) and disorganized actin filament structures in endocytic mutants (340).

FIG. 3.
Electron micrographs of cortical actin patches. (A) Thin sectioning and electron microscopy reveal membrane invaginations at the yeast cell cortex that are enriched for actin (immunolabeled with 10-nm gold particles; arrow) and Abp1 (immunolabeled with ...

Using complementary EM techniques, a recent study by Rodal et al. has provided the first three-dimensional information on patch ultrastructure (314). Intact samples of the yeast cell cortex, prepared using quick freezing and deep etching, revealed densely branched networks of actin filaments labeled with antibodies to actin, Abp1, the Arp2/3 complex, and Crn1 (314) (Fig. 3C and D). The caveat to this methodology is that it requires the use of spheroplasted cells. However, spheroplasted cells maintain endocytosis (299), suggesting that the structures preserved and visualized by EM represent functional actin patches. The structures are cone shaped, with the Arp2/3 complex positioned at the apex, and there may be some relationship between these actin mounds and the finger-like invaginations reported by thin sectioning (Fig. (Fig.3).3). In another study, Young et al. (423) used partial cell lysis to leach semi-intact actin patches from cells, purify them through multiple steps, and image them by EM. The actin filaments in these structures form a branched network, and decoration with myosin S1 fragment showed that the filaments have uniform polarity. When combined with the in vivo observations of Rodal et al. (316) showing that the Arp2/3 complex (which binds to filament pointed ends) is positioned at the apex of cone-shaped actin patches, these observations suggest that the majority of actin filament barbed ends in actin patches may be oriented towards the cell cortex. This assignment of filament polarity also is supported by photobleaching experiments in sla2 mutants (173), which block patch internalization and lead to hyperextended patch-like structures. A photobleached band within this extended structure moves away from the cell cortex, suggesting that new monomers are incorporated on barbed ends of filaments at the cortex.

Unified model for actin patch development and function.

In this section, we provide a working model for actin patch development and endocytosis, based on the work described above and additional studies introduced below (Fig. (Fig.44).

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 ...

(i) Early recruitment: nonmotile phase.

The early recruitment process may begin with cytosolic regions of membrane receptors, possibly after being modified by kinases and ubiquitin ligases, associating with endocytic machinery (147, 156, 323, 346, 362). Alternatively, some of the early endocytic machinery may reside in patches at the cell cortex and recruit and concentrate receptors. Supporting this second possibility, a recent study has described the existence of a static complex—termed the eisosome—that marks the future site of actin patch formation and contains the proteins Pil1, Lsp1, and Sur7 (390). It is important to note that actin patches also formed at sites independent of eisosomes, suggesting multiple mechanisms for specifying sites of actin patch assembly. The initial endocytic machinery recruited to patches includes clathrin and multiple scaffolds and clathrin adaptors: Yap1801 and Yap1802 (AP180 homologues), Ent1 and Ent2 (epsin homologues), Ede1 (Eps15R homologue), Scd5, Sla1, and Sla2 (6, 115, 145, 156, 266). These proteins recruit additional early patch components that promote actin assembly, such as Pan1, End3, and Las17, to form a complex network of interacting factors. Specifically, the nucleation-promoting factors (NPFs) (see below) Pan1 and Las17 may directly recruit and activate the Arp2/3 complex to nucleate actin assembly.

Of the many proteins involved in this early step of patch formation, two well-studied components that seem to play central roles are Sla1 and Sla2. Sla1 binds directly to receptors to promote their internalization by a ubiquitin-independent mechanism (156). Sla1 also appears to be important for recruiting other factors to patches, as suggested by sla1Δ mutations causing delayed initiation of actin polymerization (i.e., stalling in stage 1). Sla1 interacts with Pan1, End3 (364), Las17 (213), and Sla2 (131). Sla1 has been shown to recruit Sla2 to patches (131). Sla2 localization to patches also requires Scd5, an essential protein that interacts with early patch components and regulates Glc7 phosphatase activity (145).

Sla2 is a multidomain protein that makes numerous contributions to patch development and function. First, Sla2 may help 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 with clathrin in vivo and directly promoting assembly of clathrin in vitro (94, 95), and (iii) two-hybrid interactions between clathrin and the Sla2 coiled-coil domain (144). Second, Sla2 may bind to and regulate Rvs167, the yeast homologue of amphiphysin. The BAR domains of amphiphysin/Rvs proteins promote membrane curvature to facilitate vesicle budding in endocytosis (291). Third, Sla2 binds directly to filamentous actin via its carboxyl-terminal talin-like domain (239, 240). Although deletion of this domain from Sla2 causes no obvious defects in endocytosis (403, 420), this fragment of Sla2 is required for cell growth and endocytosis in the absence of specific clathrin adaptors, suggesting an important overlapping role in endocytosis (20). Thus, it will be interesting to learn if and how Sla2 affects actin dynamics and organization. Fourth, through its amino-terminal AP180 N-terminal homology domain, Sla2 binds to phosphoinositide PI(4,5)P (PIP2) to facilitate the internalization step of receptor-mediated endocytosis (360). Finally, as Sla2 binds to Sla1 (18), which in turn binds two NPFs (Pan1 and Las17), it is possible that Sla2 also contributes to the spatial and temporal regulation of Arp2/3 complex activity.

(ii) Intermediate stage in patch development: slow motility.

Transition to the second stage in patch maturation, which likely is coupled to membrane invagination, occurs when early patches (marked by Las17, Pan1, Sla1, and Sla2) are joined by the actin nucleation machinery, consisting of the Arp2/3 complex and three new NPFs, Abp1, Myo3, and Myo5 (type I myosins). Coincident with the detection of filamentous actin, patches begin to undergo slow, nondirectional movements within the plane of the cortex. These movements are sensitive to Lat-A, suggesting that the transition to this intermediate stage is driven by actin polymerization (173). Using mutants stalled at this stage of patch development, the measured rates of actin polymerization were found to be 0.05 to 0.1 μm/s (173, 174), markedly slower than subsequent cable-dependent rapid patch movements (0.3 μm/s) (159).

(iii) Scission and rapid transport of patches.

Transition to the third phase of patch development correlates with the onset of rapid, directional patch motility, in which patches/vesicles move inward from the cell cortex (173) and simultaneously shed many of their early components (e.g., Sla1, Sla2, Las17, Pan1, Myo3, and Myo5). The signals that trigger the transition from slow-moving to fast-moving patches are unknown, but based on several lines of evidence, Ark1/Prk1 kinases are implicated. These kinases associate directly with Abp1 and Sla2 to localize to patches and directly phosphorylate Sla1 and Pan1, and possibly other patch components (reviewed in reference 355). Phosphorylation of Pan1 by Prk1 inhibits its ability to bind actin and activate the Arp2/3 complex (371). Further, chemically induced loss of Ark1/Prk1 kinase activity in vivo using analog-sensitive kinase mutants causes the rapid and Arp2/3-dependent formation of actin clumps (340). Actin clumps consist of endocytic vesicles that have failed to mature properly and are decorated with Abp1p, Sla2p, Pan1p, Sla1p, and Ent1p. Therefore, Ark1 and Prk1 may negatively regulate the actin assembly-stimulating activity of early endocytic proteins and thereby play a critical role in directing vesicle internalization 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 these events. Las17 is shed prior to Pan1 (173), suggesting that Las17 and Pan1 have different roles in vesicle internalization. In addition, mutations in Bbc1, End3, Sla1, and Sla2 (alone and in combinations) show formation of dramatic actin protrusions (174), suggesting that they may contribute to the down-regulation of actin assembly at this stage and/or help promote vesicle release. Consistent with this view, purified Sla1 and Bbc1 directly bind to and inhibit the NPF activity of Las17, and sla1Δ and sla2Δ cells show defects in patch ultrastructure by EM (314, 315). Deletion of SLA1 results in large, flattened actin patches, and deletion of SLA2 results in large, raised actin patches (314). Proper release of vesicles also requires Myo3, Myo5, Rvs161, and Rvs167, which are implicated in promoting membrane scission (172, 174). Rvs161 and Rvs167 mutants in particular show a striking phenotype where internalizing vesicles are retracted back to the cortex, suggesting a direct role in scission (174).

Once patches/vesicles leave the cell cortex, they move rapidly inward along polarized actin cables, structures described in detail below. The retrograde (bud-to-mother) flow of cables appears to “carry” patches to endosomal sorting compartments (159). The rate of patch transport is similar to the rate of cable flow, suggesting that transport is passive and must involve a physical link between patches and cables (159). The only factors known to associate with fast-moving patches are Abp1, the Arp2/3 complex, Cap1/Cap2, and Sac6 (159, 173). Whether other known F-actin-associated patch proteins and their ligands (e.g., Cof1, Crn1, Srv2, Scp1, Twf1, and Abp140) are also present in these patches remains to be determined. Of these factors, Abp140 and Sac6 are strong candidates for providing linkage between patches and cables, since both are known to decorate patches and cables (3, 17). Further, both of these proteins can cross-link actin filaments in vitro (3, 17), an activity well tailored for bridging two filamentous actin structures.

Finally, some late endosome movements have been reported to require ongoing actin polymerization at the endosome surface. These movements were dependent on both the NPF activity of Las17 and Lsb6 (a Las17-binding protein) (58, 59), and the authors suggested that Arp2/3-dependent actin assembly powers late endosome movements similar to the mechanism employed by Listeria (see above). However, one dilemma with these observations is that Las17 and actin have not been detected on late endosomes, which the authors suggest may be due to their low abundance on such structures. Thus, it remains open whether these actin-dependent movements represent a second, independent class of late endosomes, or instead the defects in late endosome motility in las17 mutants arise from aberrant assembly of endosomes at an earlier stage.

Actin Cables: Dynamic Tracks for Polarized Growth


Unlike animal cells, which rely primarily on microtubule-based transport to establish and maintain cell polarity (353), yeast cells use actin-based transport along cables to direct polarized cell growth and to segregate organelles prior to cell division (45). Early in G1, unbudded cells detect cortical landmarks remaining from the previous cell division to select a future bud site, and polarity factors (including the polarity cap; see below) are recruited to this end of the cell. From this site, actin cable assembly is initiated, which leads to reorientation of actin cables and thus targeting of growth and secretion to the future bud tip. Polarized growth towards the bud tip (or cap) continues through a medium-budded stage and depends on actin cables emanating from the bud tip and neck. 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 polarized cell growth was made possible by using a fast-acting temperature-sensitive tpm1ts tpm2Δ mutant strain (306). In this work, Pruyne and coworkers showed a complete loss of cables in tpm1ts tpm2Δ cells after a 1-minute incubation at the restrictive temperature. Cells also rapidly lost the accumulation of a secretory marker (Sec4) and a class V myosin transporter (Myo2) at the bud tip. Remarkably, cables reassembled within 1 minute after their return to the permissive temperature, and polarity markers were restored at the bud tip shortly thereafter. Similar phenotypes were observed for the temperature-sensitive myo2-66 allele, which carries a mutation in its motor domain (45, 171, 334). From an examination of the literature, it is evident that mutations in most factors known to influence cable assembly and stability lead to 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 diverse cargos along actin cables to facilitate polarized growth. Post-Golgi secretory vesicles marked by GFP-Sec4 can be imaged in real time moving directionally on cables towards the bud tip in a Myo2-dependent manner (335). These vesicles carry enzymes that contribute to the production of new cell wall required for bud growth and cell division (1, 343), including glucan synthase, which localizes to actin patches (376). In addition, organelles such as the vacuole, Golgi, nucleus, cortical endoplasmic reticulum, and peroxisomes (305, 421) are delivered to the daughter cell in a cable- and Myo2-dependent manner, and daughter-specific mRNAs (e.g., ASH1) are retained (and possibly transported) in the daughter cell in an actin cable- and Myo4-dependent manner (40, 72). Finally, mitochondria also appear to be transported along cables, and their anterograde movements occur through a myosin-independent mechanism that relies on Arp2/3 complex-based actin polymerization, reminiscent of Listeria motility (104).

Mechanism of actin cable formation.

Whereas patch formation relies on actin nucleation by the Arp2/3 complex, cables appear to assemble in an Arp2/3-independent manner (99, 407). Until recent years, it had been speculated that cables might be assembled by a mechanism of filament capture and incorporation, similar to what was proposed for the assembly of actomyosin rings and the bundled actin structures in Drosophila melanogaster bristles. However, recent studies show that cables are assembled by the actin-nucleating activity of formins and profilin. Two key observations led to this discovery. First, it was shown that the formins Bni1 and Bnr1 are acutely required for cable assembly (99, 325), consistent with their localization to sites from which cables emanate (182, 284, 304) (Fig. (Fig.5A).5A). When conditional bni1ts bnr1Δ mutant cells were shifted to the nonpermissive temperature, all visible actin cables disappeared within 2 min (Fig. (Fig.5B),5B), and cables reappeared within 2 to 4 min upon return to the permissive temperature (99, 325). Second, biochemical analyses demonstrated that purified carboxyl-terminal fragments of Bni1 directly nucleate actin assembly in vitro (303, 326). These papers inspired studies on formin activity in other species, and it is now evident that actin assembly-promoting activity is a universal property and function of all formins examined in animals, plants, and fungi (149, 194).

FIG. 5.
Formins are required for actin cable assembly at the bud tip and neck. (A) Localization of GFP-labeled yeast formins Bni1 and Bnr1 throughout the cell cycle. (Reprinted from reference 304 with permission of the publisher.) (B) Formin function is required ...

Despite these advances, key questions remain regarding cable formation. How are the potent actin nucleation activities of Bni1 and Bnr1 regulated? Studies in mammalian cells suggest that formins are auto-inhibited until released by binding of activated Rho family GTPases to the formin Rho-binding domain (389); however, it has not been determined if Bni1 and/or Bnr1 are auto-inhibited. In yeast, there is evidence to suggest that Cdc42 contributes to formin regulation during early bud emergence and then the partially redundant Rho3 and Rho4 regulate formins during most phases of cell growth, whereas Rho1 may regulate formins in response to cell stresses (79). The specific molecular mechanism underlying this in vivo control remains to be defined. Do Cdc42, Rho1, Rho3, and Rho4 all act by binding to Bni1 and Bnr1 to relieve a proposed auto-inhibited state? Do they act specifically on Bni1 or Bnr1? Are other factors required for activating formins? How are Bni1 and Bnr1 functions regulated by phosphorylation (122, 235)? Once nucleated, how are actin filaments spatially organized into cables? Is this mediated by the proteins that decorate cables, such as Tpm1, Tpm2, Sac6, and Abp140 (17, 84, 219, 306)?

The bud tip polarity cap.

The term “polarity cap” refers to a group of interacting cellular factors that localize primarily to the bud tip during bud emergence and growth and have genetic roles in directing polarized cell growth. Subsequently, many of the polarity cap components shift localization to the bud neck, just prior to cell division, where they appear to facilitate cytokinesis by mechanisms described in the next section. Some polarity cap components help to promote cable assembly, as discussed below, while others regulate dynamic membrane trafficking events (exocytosis and endocytosis) and additional aspects of cell polarity at the bud tip; for more details see other recent reviews (54, 138, 157, 302).

One functional complex within the polarity cap network has been referred to as the polarisome. Its three components—Spa2, Pea2, and Bud6—comigrate as part of a 12S complex in cell extracts fractionated by sedimentation velocity (345). In addition, the Rho family GTPase Cdc42 and two Cdc42 effectors, Bni1 and Gic2, appear to function intimately with the polarisome, as supported by numerous two-hybrid and coimmunoprecipitation interactions with Spa2, Pea2, and Bud6 (97, 113, 169). Dozens of other components now are routinely referred to as part of the polarity cap network based on their physical interactions and localization pattern to the bud tip.

The concept of a large macromolecular assemblage directing polarized cell growth raises a number of interesting questions. First, what are all the components? To date, at least 60 proteins have been identified that localize to the bud tip, most of which show physical and/or genetic interactions that suggest their involvement in regulating polarity (165). Undoubtedly, this list will continue to grow. Second, are there stable subcomplexes within the larger assemblage? As mentioned above, at least one stable complex has been identified, containing Spa2, Pea2, and Bud6, and there is evidence that this same complex may contain Gic2 (169) and possibly the Spa2-related protein Sph1 (16, 319). Greater efforts are required now to isolate biochemically and define by mass spectrometry the components of this and other bud tip complexes. Third, how are the interactions among complexes and components spatially and temporally regulated? As has proven to be the case for other large biological machines (e.g., kinetochore, centrosome, and spliceosome), there are likely to be stable subcomplexes as well as dynamically interacting components. Specific components may change interactions in response to molecular signals. Two kinases that regulate cell polarity are implicated in phosphorylating Bni1, a central figure in the polarisome because of its role in nucleating actin assembly. Bni1 is phosphorylated directly by Fus3 kinase and is phosphorylated in a Ste20 kinase-dependent manner in vivo (122, 235). The Bni1-interacting protein Bud6 is phosphorylated (251); further, it associates with Ste11 (a MEK kinase), which regulates polarity (251, 345). In addition, Bni1, Gic2, and the PAK-like kinases Ste20 and Cla4 are directly regulated by interactions with Rho GTPases (169, 170). These provide inspiring leads into the mechanisms regulating polarity cap assembly and function but also likely represent only a small percentage of the total signaling events involved.

A more daunting question, and one that will likely require combined 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 cables, and (iii) receiving positive feedback signals that help sustain polarity. Several polarity cap components (Bud6, Cdc42, Rho3, Rho4, and profilin) are thought to regulate Bni1 directly to promote actin cable assembly (see below). In addition, Spa2 directly interacts with Bni1 and Pea2 (113, 345), and Spa2 and Pea2 are required for Bni1 localization (284). The biochemical 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 in early bud emergence (169), and gic1 gic2 mutants show defects in establishment of polarity early in bud emergence (47, 60). Since Bud6 regulates Bni1 activity (253), it will be interesting to 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 a working model for the polarity cap, focused on its role in promoting actin cable assembly (Fig. (Fig.6).6). Future models will need to incorporate mechanisms for secretory vesicle docking and fusion, other membrane remodeling events, and cell wall synthesis. In addition, a clear picture of polarity cap architecture and function eventually will require defining (i) the relative abundances of each component, (ii) the stable protein complexes formed among components, (iii) the rules for hierarchical assembly and localization of components at the bud tip, and (iv) functional interactions linking different nodes of the 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 and cellular morphogenesis] signaling, and mitogen-activated protein kinase signaling) (165). For instance, Msb3 and Msb4 appear to coordinate multiple functions in polarized cell growth, including regulating the activity of the GTPase Sec4 in exocytosis, binding to the central polarity cap factor Spa2, and controlling Cdc42 activity (366).

FIG. 6.
Models for regulation of actin cable assembly at the bud tip and neck. (A) Organization of formins (red), actin cables (gold), and septin structures (blue) in a budded yeast cell. Formin proteins (Bni1 at the bud tip and Bnr1 at the neck) directly nucleate ...

Actin cable architecture.

Cables must provide polarized tracks for continuous myosin-dependent transport of cargos to the bud neck and tip. Therefore, early models suggested that cables might be stable structures, comprised of long bundled actin filaments, providing relatively stationary tracks for myosin movement. However, it is now evident that cables are comprised of shorter filaments organized into bundles of uniform polarity, with the majority of their fast-growing (barbed) ends oriented towards polarity sites. Evidence for this organization includes the following: (i) a single fluorescently labeled cable has variations in its width, suggesting the 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) myosin V-dependent transport is unidirectional towards polarity sites (335), and yeast type V myosins (Myo2 and Myo4) are barbed end-directed motors (310); and finally, (iv) formins associate with the barbed ends of actin filaments and localize to the bud tip and neck. The precise arrangement of actin filaments in S. cerevisiae cables remains uncertain, because cables traverse multiple planes within a cell and have eluded description by thin-section EM and tomography. However, the recent use of permeabilized cells for EM in S. pombe has revealed that actin cables are comprised of short overlapping filaments with most of the barbed ends oriented towards the cell tips, consistent with facilitation of barbed end-directed myosin movement (175). Given all of the points above, it seems likely that S. cerevisiae cables will show a similar organization.

Gone in 60 seconds: actin cables are composed of short filaments.

Intriguingly, actin cables are lost within 60 seconds of treating cells with the actin monomer-sequestering drug Lat-A (19). This finding demonstrates two important properties. First, the filaments in cables likely undergo rapid turnover, as Lat-A sequesters monomers to inhibit new actin assembly but is not reported to promote filament disassembly. Second, due to the rapid nature of cable loss, the filaments in cables must be relatively short. In fact, their approximate length can be estimated from their disassembly rate using the following simple calculations. A typical large-budded yeast cell is ∼10 μm long, with actin cables ∼4 μm long. Each actin subunit within a filament contributes 2.77 nm to filament length (155), so if this 4-μm cable is comprised of a single linear filament, it contains ∼1,500 subunits. The barbed end of this filament may be capped by formins (see below), but to make a more liberal estimate of filament length we will assume that both filament ends are uncapped. Using the dissociation rates for rabbit skeletal 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 barbed end, 0.8 s−1 for the pointed end) (293), a 4-μm filament (1,500 subunits long) would lose about 450 subunits in 1 minute if composed entirely of ADP-actin or 130 subunits for all ATP-actin. In either case, a 4-μm-long filament in a cable would maintain 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 filament length in cables is <1 μm. These estimations do not consider the stabilizing and destabilizing effects of actin-binding proteins that decorate the sides of cables. Nonetheless, these estimates for S. cerevisiae are remarkably consistent with the measured lengths of filaments (0.4 to 0.5 μm) in S. pombe cables as determined by EM (175).

This still leaves many open questions concerning cable regulation. How is the length of filaments in cables controlled? The extent of filament elongation could be restricted by capping proteins. While no conventional barbed or pointed end capping proteins have been visualized on cables, it is possible their decoration is sparse (since they associate only with filament ends) and thus has escaped detection. Cap1/Cap2 (yeast capping protein), which associates tightly with filament barbed ends to block both addition and dissociation of subunits, is a strong candidate to be on cables, because cap2Δ cells exhibit dramatically reduced cables (10). This phenotype is consistent with Cap1/Cap2 having a role in stabilizing the filaments in cables. Formins also bind tightly to barbed ends but permit filament growth through a processive capping mechanism and protect growing ends from Cap1/Cap2 (253, 429). Thus, formins may initiate actin nucleation at the bud tip but remain associated with filaments as they are incorporated into cables. Consistent with this model, actin cables undergo retrograde flow away from the cortical sites of polarized growth (see below), and formin-GFP punctae have been visualized on moving actin cables in S. cerevisiae and S. pombe (D. Pellman and F. Chang, personal communications). Thus, the capped state of filaments in cables is an important issue to resolve and raises even more questions: are formins present on all or a subset of filaments 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 in light 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 perform this function in vertebrate muscle and nonmuscle cells, tightly capping the pointed ends of tropomyosin-decorated filaments (108). Although obvious tropomodulin homologues have not been identified in the S. cerevisiae genome, functional homologues may exist, mutants of which likely would cause diminished cables.

Actin cable dynamics and turnover.

The observed rapid turnover of cables suggests that there may be active mechanisms required for disassembling the filaments comprising cables. However, in wild-type cells, the primary filament disassembly factor, cofilin, is not detected on cables. Instead, cables are decorated with tropomyosin, which stabilizes filaments and competes with cofilin for binding F-actin (30, 278). These observations have left the high rate of cable turnover unexplained; however, two studies provide insights into possible turnover mechanisms. First, cofilin was shown to decorate cables in aip1Δ cells (317). Second, cofilin and Aip1 were demonstrated to promote rapid cable turnover (276), with cof1-22 and aip1Δ mutations causing ∼20- and 5-fold decreases in the rate of cable turnover, respectively. This work suggests that most filaments in cables are decorated and stabilized by tropomyosins Tpm1 and Tpm2, but subsets of filaments are stochastically and rapidly disassembled by the combined activities of cofilin and Aip1 (see “Rapid Turnover of Actin Structures” for more details). This leads to a steady thinning or “pruning” of cables along their lengths. Given the discovery of this function for cofilin and Aip1, other factors known to promote filament disassembly and turnover should be tested for their roles in cable turnover (e.g., Srv2/CAP, profilin, and twinfilin).

Using Abp140-GFP as a marker, Yang and Pon have imaged actin cable dynamics in real time in live cells (419). Their work shows that for many cables, one end associates with the bud tip or bud neck while the other end moves away from these assembly sites (in the direction of the mother cell) at ∼0.3 μm/s. Some cables appear to assemble at the bud tip and extend through the bud neck into the mother compartment. Other cables appear to move along the cell cortex without having one end attached to the bud tip or bud neck. It is not yet clear whether these “free-roaming” cables have distinct functions and whether they are initially assembled at the bud tip and neck and detach from those sites, or instead form independent of the polarity sites. The direction of cable flow (retrograde, away from the bud tip) is somewhat unexpected given that cables serve as tracks for myosin V-based barbed end-directed transport in the opposite direction (anterograde, toward the bud neck and tip). However, retrograde cable flow (0.3 μm/s) does not appear to pose a major difficulty to overcome, since myosin V-dependent anterograde transport is about 10 times faster—4.5 μm/s in vitro (310) and 3 μm/s in vivo (335).

What provides the force that drives cables away from the bud tip and neck and toward the mother cell? One possible answer is the force generated by actin polymerization. As formins polymerize actin filaments at the bud tip and neck, these elongating filaments may be pushed into the mother cell due to insertional polymerization at their barbed ends capped by Bni1 and Bnr1 (197, 251, 253, 429). If cable flow rate (0.3 μm/s) were directly proportional to actin polymerization rate, this would require filament elongation rates of ∼100 subunits/s. This would require 5- to 10-μM monomeric actin in the cytosol, which is unlikely to exist. However, interactions between profilin, actin monomers, and the formins Bni1 and Bnr1 may accelerate cable assembly rates to match rapid cable flow rates (see “Profilin-FH1 interactions: a throttle for filament elongation” below).

In addition, cable flow rates are likely influenced by cellular mechanisms beyond actin polymerization, including a mechanism recently suggested to involve myosin. Domain and sequence similarities among myosin heavy chains define myosin classes, also called types (200, 400). S. cerevisiae expresses five myosins: two type V myosins (Myo2 and Myo4) that transport secretory vesicles, organelles, and mRNAs to sites of polarized growth (bud tip and neck); a single type II myosin (Myo1) that promotes formation and closure of the cytokinetic ring; and two type I myosins (Myo3 and Myo5) that promote actin assembly and endocytosis at cortical patches (22, 126, 305). Recent work suggests that Myo1 has an important role in controlling actin cable dynamics (T. Huckaba and L. Pon, personal communication). Myo1 first localizes to the incipient bud site early in G1 (33, 204, 362) and then is found at the bud neck from the earliest stages of bud formation. Although Myo1 is thought to function primarily in cytokinesis, much later in the cell cycle, Huckaba and colleagues observe reduced cable velocities in the absence of Myo1 motor activity. They propose that Myo1, situated at the bud neck, uses its barbed end-directed motor to “pull” cables through the bud neck and facilitate directional flow toward the mother cell. Thus, cable movement may be regulated by actin polymerization, Myo1 activity, and possibly other factors.

Yeast Cytokinesis: Two Complementary Mechanisms for Cell Division


Virtually all animals and fungi utilize an actin-based contractile ring to physically separate the two daughter cells during cytokinesis. Although a cytokinetic actin ring had been observed in fission yeast as early as 1986 (231), the existence of such a division mechanism in budding yeast was realized only in recent years. Early work suggested an enrichment of actin at the site of cell division in yeast (2, 183), but an actomyosin contractile ring was not observed until 1998 (34, 217). In animal cells, myosin II controls contraction of the actin cytokinetic ring (105), which constricts the cleavage furrow to a 1- to 2-μm-wide midbody zone (258, 308, 327). In S. cerevisiae, the bud neck is less than 1 μm throughout the cell cycle; this led to the suggestion that constriction would not occur in yeast and, instead, that targeting of secretory vesicles for membrane and cell wall deposition to the bud neck would be sufficient to drive cell division (327). This view was reinforced by the 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 structural role rather than a dynamic role at the bud neck. Further, since some yeast strains lacking MYO1 were shown to be viable and complete cytokinesis, its role in cytokinesis remained dubious (34, 318, 395). Then, in 1998, two seminal studies imaged Myo1-GFP dynamics in live cells and provided clear evidence that a myosin ring contracts specifically during cytokinesis (Fig. (Fig.7)7) (34, 217).

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 ...

It is now evident that two complementary mechanisms contribute to yeast cytokinesis: first, contraction of an actomyosin ring, and second, formation of a septum, achieved by delivery of secretory vesicles to the bud neck to facilitate membrane deposition and cell wall synthesis. It remains unclear how these two mechanisms are coordinated, as discussed below.

The position of actin ring assembly (and the future plane of cell division) in budding yeast is dictated by the position of bud emergence initiated in early G1, where the ring will be assembled at the neck of the newly growing bud (54, 302, 305). The bud site is selected by cortical cues (bud site selection markers) remaining from the previous cell division. Early in G1, these positional cues trigger recruitment and activation of a cascade of factors essential for assembling both the polarity cap at the incipient bud tip and components of the bud neck. At the top of this hierarchy of bud neck components are the septins, which form a ring structure discussed in detail below. Through poorly understood mechanisms, this scaffold in turn recruits Myo1, formins Bni1 and Bnr1, two myosin light chains (Mlc1 and Mlc2), Hof1/Cyk2, IQGAP (Iqg1/Cyk1), and Cyk3 (22). All of these factors arrive at the bud neck before visible actin staining with the exception of Cyk3, which appears during anaphase.

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

These observations raise many important questions, including the following. What are the signals at anaphase that induce cable reorientation and actin ring assembly at the neck? What is the architecture of filaments in the ring? What are the signals that trigger ring contraction? How are the activities of cables and the ring coordinated to facilitate cell division? Each of these issues is discussed in greater detail in the following sections.

Targeting secretion to the bud neck at cytokinesis.

Prior to anaphase, polarized actin cables are assembled at two locations, the bud tip (where Bni1 localizes) and the bud neck (where Bnr1 localizes). This produces two sets of cables that together target secretion to the bud tip. At anaphase, Bni1 abruptly changes localization, joining Bnr1 at the bud neck (176, 284, 304). In a similar time frame, other components of the polarity cap, such as Spa2 and Bud6, change localization from the bud tip to the bud neck (14, 345). These events redirect secretion to the bud neck, as indicated by relocation to the neck of Myo2, Mlc1, and exocytosis markers such as Sec3 and components of the exocyst complex (305). Targeted delivery of new plasma membrane and cell wall materials provides additional surface area required for cell division. Other key factors delivered to the neck include Chs2 and Chs3, the catalytic subunits of chitin synthase II and chitin synthase III. Chs2 contributes to formation of the primary septum, while Chs3 functions primarily in bud scar chitin synthesis; however, these two enzymes appear to share an essential role in septum formation (63, 343). chs2Δ and chs3Δ cells are viable but show defects in septum formation and cell division, and chs2Δ chs3Δ double mutants are inviable and arrest at cytokinesis (343). This demonstrates an essential role for targeted septum deposition and cell wall synthesis in cytokinesis. Remarkably, these secretion-driven pathways are sufficient for completing cytokinesis in the absence of a contractile actin ring, albeit not as efficiently, suggesting that they make the dominant contribution to cell division.

Formation and closure of the actomyosin ring.

Actin filaments are among the final components to be assembled into the contractile ring and do not become visible until late anaphase. 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 the bud neck in order to form the ring (105). However, more- recent evidence suggests that ongoing actin assembly by Bni1 and Bnr1 is required for ring formation and function. While the actin ring forms in either bni1Δ or bnr1Δ single mutants (378), a temperature-sensitive formin mutant strain (bni1ts bni1Δ) fails to assemble the ring (369). Other factors required for ring assembly include profilin and tropomyosin (369). Thus, the machinery driving ring formation is highly similar to that driving cable assembly. In S. pombe, formins are essential for actin ring formation and cytokinesis (57), but the Arp2/3 complex also localizes to the actomyosin ring and makes a contribution to its formation (288), possibly by promoting endocytosis to facilitate septum formation. In contrast, actin ring formation in S. cerevisiae is thought to occur independently of the Arp2/3 complex, and to date the Arp2/3 complex has not been localized to the actin ring. Despite this prevailing model, two regulators of the Arp2/3 complex, Las17 and Vrp1, have known roles in cytokinesis (265, 367), and actin structures that depend on Arp2/3 complex activity have been observed at the bud neck (270), leaving open many possibilities for the mechanism of their involvement.

Three questions regarding the behavior of the assembled actin ring point to important issues for future investigation. First, how are actin filaments in the ring organized? A requirement for actin polymerization by formins suggests that filaments should be unbranched. Further, the classic “purse string” model for ring contraction predicts a network of filaments with antiparallel arrangement to facilitate myosin-based contraction (109). Resolving this will require ultrastructural analysis of partially purified actomyosin rings and/or intact rings in cells. Second, what is the mechanism driving actin ring contraction? One possibility is the motor activity of yeast type II myosin (Myo1). Indeed, a point mutation impairing F-actin binding 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 detectable defects in cytokinesis (224). While these data appear to be at odds, perhaps instead they suggest that the Myo1 motor domain contributes to force production during cytokinesis but can be replaced by other cellular factors (possibly other myosins) when deleted. Another possibility is that nonmotor activities of Myo1 promote ring closure. Recent work in mammalian cells suggests that type II myosin promotes actomyosin ring disassembly rather than contraction (133, 263). Myo1 may employ a similar mechanism to facilitate cytokinesis, but this model awaits testing. It will also be important to identify the signals that regulate Myo1 activity (and thus the timing of ring contraction). Third, how are actin filaments in the dynamic actomyosin ring disassembled? Myo1 may contribute to disassembly, as discussed above, but additional factors are likely involved; one example is cofilin, which functions at the cytokinetic ring in S. pombe (264). Clearly, we have much to learn about how the many ring components that promote actin filament assembly (e.g., formins, profilin), stabilization (e.g., tropomyosin, Sac6), and turnover (e.g., cofilin) are exquisitely regulated to control ring formation and contraction.

Role of yeast myosin light chains in cytokinesis.

All myosins are comprised of both heavy and light chains. The heavy chains are large polypeptides (typically 100 to 200 kDa) with a motor domain, multiple IQ motifs, and variable tail regions. The light chains are small polypeptides (∼15 kDa) with a calmodulin-like fold that bind to IQ motifs in heavy chains to facilitate motor activity. Further, calcium binding and/or phosphorylation of light chains can regulate their association with heavy chains, providing cells with a mechanism for controlling myosin activity. 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 their roles 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). Differences in the functions of the light chains are apparent from their distinct patterns of localization and mutant phenotypes. Mlc2 colocalizes with Myo1 at the bud neck and requires Myo1 for localization (227). In contrast, Mlc1 localizes to the bud neck independently of Myo1 (342). Deletion of MLC2 has little effect on cell growth or cytokinesis but causes a slight delay in Myo1 ring disassembly (227). In contrast, MLC1 is essential for cytokinesis and cell viability (33). Since myo1Δ cells are viable and can undergo cytokinesis, the lethality of mlc1Δ suggests that it performs additional functions beyond regulating Myo1. This is further suggested by disruption of Mlc1-Myo1 interactions with specific point mutations in Mlc1 or deletion of the IQ motifs in Myo1, both of which cause 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 that promote cytokinesis, Myo2 (358) and Iqg1/Cyk1 (IQGAP) (342). Its interaction with Myo2 targets vesicles to fill the bud neck, a key step in formation of the septum (388). However, removal of all six IQ motifs from Myo2 (i.e., the myo2-Δ6IQ mutant) does not affect growth or division rates (358), indicating that the Mlc1-Myo2 interaction is not essential for motor function and likely serves primarily in a regulatory capacity. Mlc1 interactions with Iqg1 may have multiple roles. Iqg1 requires Mlc1 for its localization to and function in assembling the cytokinetic ring (96, 341). Iqg1 also has been implicated in promoting septum formation (280), which could require its interactions with Mlc1 and is consistent with the IQG1 gene being essential (96). Thus, Mlc1 appears to coordinate both actin-based pathways facilitating cytokinesis: (i) polarized transport of vesicles to form the septum and (ii) formation and closure of the actomyosin ring. One key open question is how Mlc1 localization is regulated, since there are no known ligands 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 vesicle delivery and ring assembly and closure) which are genetically separable and are regulated by a large number of proteins found at the bud neck (121). The temporal sensitivity and complexity of these two pathways suggest that they must be coordinated; however, the mechanisms for this remain poorly understood. One family of proteins that may be involved is the formins (Bni1 and Bnr1), as they directly nucleate the actin structures 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 for actin ring formation, and the requirement for Rho1 is overcome by expression of a dominant-active Bni1 construct (369). This suggests that Rho1 functions upstream of Bni1 to stimulate actin ring assembly. Functions for other yeast Rho GTPases during cytokinesis remain untested.

Another strong candidate for coordinating septum formation with contractile ring function is Hof1/Cyk2, a member of the cdc15/PSTPIP family of proteins (216). Deletion of HOF1/CYK2 causes temperature-sensitive growth and leads to defects in both septum formation and ring contraction at the nonpermissive temperature (215, 378). Hof1 localizes to the bud neck independent of Myo1 (215, 378) and contracts only partially during cell division (unlike Myo1) (215, 378), and hof1 and myo1 deletions are synthetic lethal (378). These observations indicate that Hof1 and Myo1 perform distinct functions in cytokinesis and suggest that Hof1 may function in septum formation. Hof1 physically interacts with Bnr1, suggesting it may regulate cable assembly at the bud neck. Further, another member of the cdc15/PSTPIP protein family, S. pombe Cdc15p, interacts with both the cytokinetic formin Cdc12p and the Arp2/3-activating type I myosin (53). The genetic data implicating Hof1 in septum formation, along with physical interactions of Cdc15 and Hof1 with actin polymerization machinery, together suggest that these proteins may link and coordinate actomyosin ring formation/contraction with septum formation. These processes may be facilitated by the ability of cdc15/PSTPIP proteins to associate with and deform membranes 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., synthetic lethal with myo1) (193), and has been proposed to function with Hof1 in septum formation (33). Overexpression of either CYK3 or HOF1 suppresses the lethality of iqg1Δ (193), suggesting that CYK3 and HOF1 may act downstream or are redundant with IQG1. Further, cyk3Δ and hof1Δ mutations are synthetic lethal, suggesting that they may perform separate functions downstream of Iqg1 (193).


The molecular machinery that assembles the actomyosin contractile ring and the septum must be targeted to the bud neck. The timing of arrival and loss of different components at the bud neck varies greatly throughout the cell cycle (121), and the mechanisms for recruitment and maintenance of localization of these proteins are just beginning to emerge. Some of the earliest proteins to arrive are the septins (including Cdc3, Cdc10, Cdc11, and Cdc12 in budding yeast), which in turn are required for localization of many other proteins (222). Septins are ubiquitous GTP-binding proteins that form stable heteromeric complexes; these units assemble tandemly into 10-nm-wide filaments that can be organized further into polymer systems (106, 187). Septin filaments in cells were first observed by EM studies as striations at the bud neck that disperse in septin mutants (48, 49). Later, septins were shown by immunofluorescence to arrive at the cell cortex prior to bud emergence (110, 184) and shortly thereafter reorganize into a collar that persists at the bud neck until after the completion of cell division (167). Initial assembly of these septin structures depends on the activity of the GTPase Cdc42 (65, 119, 120, 167) and two of its downstream effectors, Gic1 and Gic2 (167). Purified yeast septins assemble into filaments in vitro (111), and GTP binding is required for their polymerization and in vivo functions (381).

Formation of the actomyosin ring depends on septin integrity (34, 217); however, only the Myo1 ring contracts (the septin ring does not), indicating that they are separate structures (110, 184, 222). In addition to positioning the actomyosin ring, septins recruit enzymes, such as chitin synthase II, required to form the septum (33). One model to explain these critical roles of septins is that they are near the top of a physical interaction network or hierarchy for assembly of components at the bud neck. However, potential physical interactions between septins and the actomyosin ring machinery have remained elusive in yeast. In animal cells, the adaptor protein anillin physically links actin and septin filaments in vitro (188). The fission yeast protein Mid1p has some sequence homology with anillin and functions in placement of the contractile ring at the middle of the cell (56). However, Mid1p has not been shown to bind to actin or septins; rather, Mid1p may interact with type II myosin at the division site (254). No clear anillin homologue exists in budding yeast, but Mid1 and anillin do share limited sequence homology with the bud site selection protein Bud4 (29, 270, 328, 365). Further, Bud4 localizes to the neck (328) and binds Iqg1, consistent with a cytokinetic function (280). So, Bud4 is a reasonable candidate for actin-septin linkage functions. In addition, budding yeast Afr1 may facilitate actin-septin interactions, as it associates with Iqg1 and the septin protein Cdc12 (191, 279). However, the role of Afr1 may be specialized for mating projection formation during the cellular response to pheromone (118).

Other recent work has defined a nonphysical anchoring mechanism by which septins maintain localization of some proteins at the bud neck. Prior to cytokinesis, the septin ring splits into two separate rings (223), and this split ring structure compartmentalizes and thereby concentrates (i.e., corrals) factors such as Spa2 and Chs2 at the bud neck (77). In wild-type cells, Spa2 diffuses freely within this small zone between the split septin rings. However, in cdc12-6 mutants, Spa2 and Chs2 rapidly diffuse away from the bud neck after cells are shifted to the nonpermissive temperature and the split septin ring organization is lost (77). Thus, septins may have dual roles in localizing cytokinetic machinery to the bud neck, serving possibly as both physical scaffolds and diffusion barriers.



Below, we describe the properties and mechanisms of the S. cerevisiae actin-binding proteins known to affect actin dynamics and/or organization, summarized in Table Table1.1. In each case, we attempt to relate the activities of these proteins back to their cellular roles. Our discussion includes the type I myosins Myo3 and Myo5, which help promote actin assembly, but not type II and V myosins, which have been reviewed elsewhere in detail (44, 236, 305, 324, 373).

Biochemical activities and cellular localization of yeast actin-binding proteins

Biochemical properties of actin.

Actin is a globular 42-kDa protein that exists in dynamic equilibrium between two states, monomer and filamentous polymer. The double-helical filaments assemble in a head-to-tail manner, imparting polarity to the filament. The two ends of the filament are referred to as barbed and pointed, owing to the arrowhead pattern observed upon decoration with the S1 fragment of myosin (296). Assembly of actin monomers into a filament involves an initial nucleation step, which is inherently slow due to instability of actin dimers and trimers. However, once assembled, filaments are stable and undergo rapid polarized growth, with monomeric ATP-actin subunits preferentially adding to the barbed end (11.6 subunits μM−1 s−1) versus the pointed end (1.3 subunits μM−1 s−1) (293). Addition of an ATP-actin subunit to the barbed end of a filament triggers hydrolysis of ATP bound to that subunit (half-life [t1/2]), ∼5 seconds) (295), producing actin bound to ADP and Pi. In a subsequent (slower) step, actin subunits release Pi (t1/2, ∼5 min) to produce actin bound to ADP (295). Thus, filaments at steady state (undergoing addition and loss at their ends) are a mosaic, enriched in ATP-bound actin near barbed ends, ADP-Pi actin in the middle, and ADP-actin at the pointed ends. ADP-actin subunits dissociate from filament pointed ends, and the resulting ADP-actin monomer pool must undergo nucleotide exchange (ATP for ADP) to “recharge” monomers for subsequent rounds of barbed end addition. This property of net subunit addition occurring at barbed ends and net subunit dissociation occurring at pointed ends leads to steady-state cycling of subunits through filaments, called treadmilling (Fig. (Fig.8)8) (268). Actin turnover refers to the collective dynamic events of actin subunits treadmilling through filaments, dissociating from filament ends and, as monomers, undergoing nucleotide exchange to recycle them for new rounds of polymerization.

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 ...

Each step in the process of actin filament assembly and turnover represents an opportunity for cellular control in remodeling the actin cytoskeleton. All eukaryotic cells contain a core set of actin-binding proteins that regulate actin nucleation, barbed end addition, pointed end dissociation, filament stability, and nucleotide exchange on monomers. For example, the Arp2/3 complex and formins bypass the inherently slow formation of actin dimers and trimers to nucleate actin polymerization (194, 401). Filaments are also severed and rapidly disassembled by factors such as actin-depolymerizing factor (ADF)/cofilin, Aip1, twinfilin, and Srv2/CAP (24, 268). Filaments can be stabilized by capping proteins and side-binding proteins. Further, a number of actin monomer-binding proteins cooperate to replenish the assembly-competent pool of actin monomers by catalyzing the conversion of ADP monomers dissociating from filament ends to ATP monomers. The activities and mechanisms of these factors are detailed in this section, with emphasis on how these factors work together rather than alone to affect actin filament dynamics and organization.

Assembly of Actin at Cortical Patches

The Arp2/3 complex.

Actin patches arise at cortical sites where no preexisting filamentous actin can be detected and, thus, require de novo actin nucleation. Of the two known actin nucleators that are conserved from yeast to mammals (the Arp2/3 complex and formins), only the Arp2/3 complex is found at patches (248). Further, only the Arp2/3 complex has been implicated genetically in patch assembly (248, 407, 408).

The Arp2/3 complex contains seven conserved subunits, including two actin-related proteins (Arp2 and Arp3) and five unique proteins (Arc40/p40, Arc35/p35, Arc19/p20, Arc18/p21, and Arc15/p15). The essential ARP2 gene was first identified in yeast based on sequence similarity to actin (337). Subsequent work showed that arp2 temperature-sensitive mutants have defects in endocytosis and actin organization (247, 248). The other six subunits of the yeast Arp2/3 complex were identified in 1997 (407), and deletion of any subunit except ARC18 causes severe growth defects or lethality (408). Further, the terminal phenotypes of these deletion strains show a severe or complete loss of cortical actin patch staining.

The Arp2/3 complex has been proposed to nucleate actin assembly by bringing together its Arp2 and Arp3 subunits to mimic the barbed end of a filament (294). This leaves the complex bound to the pointed end of the new filament with apparent nanomolar affinity (259). The complex also can bind to the sides of a preexisting (mother) actin filament and nucleate formation of a new (daughter) filament at a 70° angle, thus producing branched actin networks (9, 35, 150) (Fig. (Fig.9C).9C). Alone, the Arp2/3 complex has minimal nucleation activity and requires an activator or NPF for strong nucleation. The crystal structure of the inactive bovine Arp2/3 complex reveals a large separation between the Arp2 and Arp3 subunits (313), and recent studies using electron microscopy and fluorescence resonance energy transfer analysis have demonstrated that WASp/Las17 induces large structural rearrangements in the complex that bring Arp2 and Arp3 closer together to nucleate actin (Fig. (Fig.9)9) (124, 316). These observations in turn raise a new set of mechanistic questions. Which subunits facilitate Arp2/3 complex binding to the sides of filaments? What are the precise rearrangements of subunits upon activation? Which subunits, and in what order, relay activation from WASp to the final repositioning of Arp2 and Arp3? Answers to these questions will likely require mutagenesis of specific subunits involved, coupled with structural analyses of mutant and/or WASp-bound complexes.

FIG. 9.
The Arp2/3 complex mechanism of actin assembly. (A) Projection structures of free and ligand-bound yeast Arp2/3 complexes (each computed from 1,000 to 3,000 separate images). Electron microscopy and single-particle analyses were used to show that the ...

Another fundamental property of Arp2/3 complex function and regulation is binding and hydrolysis of ATP by the Arp2 and 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 promote debranching of filament networks by dissociating Arp2/3 complex from the pointed ends of filaments (208). Consistent with the importance of ATP binding and hydrolysis for Arp2/3 complex activity in vitro, mutations in the nucleotide-binding pocket of Arp2 and Arp3 severely impair actin patch dynamics and endocytosis in vivo (233).


The S. cerevisiae Arp2/3 complex is regulated in vivo by five NPFs: Las17, Myo3, Myo5, Pan1, and Abp1 (87, 98, 127, 207, 406). Each NPF has an acidic motif that facilitates binding to the Arp2/3 complex, possibly to the Arp3 and ARPC1/p40 subunits (Fig. 10B) (181, 199, 286, 398). In addition, Las17 binds to G-actin, whereas Myo3, Myo5, Pan1, and Abp1 bind to F-actin (Table (Table1).1). For Las17, Pan1, and Abp1, actin binding has been shown to be critical for their NPF activities (307, 315, 371). Actin monomer-binding NPFs such as Las17 are thought to present their bound actin subunit to the Arp2 surface to facilitate nucleation. Actin filament-binding NPFs are thought to recruit the Arp2/3 complex to the sides of preformed filaments to stimulate nucleation. Understanding the precise activation mechanism of each NPF represents an important challenge for future research.

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 ...

All five NPFs colocalize with the Arp2/3 complex, and the primary function ascribed to each NPF is activation of Arp2/3-mediated actin assembly. However, each NPF arrives at a different stage of patch maturation. These and other genetic data suggest that NPFs perform distinct yet genetically overlapping functions in patch development (see below), possibly acting sequentially on the Arp2/3 complex (173). Available genetic and biochemical data suggest that Las17 provides the strongest NPF activity in forming patches, with Pan1 and Myo3/Myo5 being ancillary as NPFs. Further, the weaker NPF activity of Abp1 may promote patch transitions by competitively inhibiting stronger NPFs (71). Below, we discuss what is known about the mechanism of each NPF and relate this to their functions at patches.

(i) Las17/WASp.

Las17 (also called Bee1) was the first NPF reported in yeast and was identified by its sequence homology with mammalian WASp (213). Early studies showed that las17Δ cells have defects in patch organization, and a permeabilized cell assay that measures incorporation of fluorescent actin was used to show that Las17 is required for actin assembly at cortical sites (213). Following demonstration that mammalian WASp activates the Arp2/3 complex, a homologous WA fragment of Las17 (consisting of its WH2 [W] domain and acidic [A] motif) was shown to activate the yeast Arp2/3 complex (406). From this work, two key questions emerged regarding Las17 NPF function. First, how does Las17 activate the Arp2/3 complex? Second, how is Las17 itself regulated? Insights into both of these questions came after purification and biochemical characterization of intact, full-length Las17 (315). It was revealed that full-length Las17 has an NPF activity far more robust than that of its WA fragment, suggesting that sequences N terminal to WA make major contributions to activation of the Arp2/3 complex. This is also consistent with genetic results showing that complete deletion of LAS17 causes much more severe defects than deletion of the WA fragment alone (las17ΔWA) (406). Increased NPF activity for full-length Las17 has been postulated to arise from N-terminal regions binding to F-actin and/or the Arp2/3 complex.

WASp family members (including SCAR/WAVE, which is not conserved in yeast) can be regulated by one of two basic mechanisms, auto-inhibition (in which the N and C termini associate directly to inhibit NPF activity) or transinhibition (mediated by direct interactions of additional proteins) (43). In both mechanisms, inhibition is relieved by signals from activated Rho GTPases. For example, Cdc42 binds directly to the GTPase-binding domain (GBD) of N-WASp to relieve auto-inhibition (320), and Rac binds directly to one of the components of a proposed WAVE1 transinhibitory complex to release active WAVE1 (89), although this mechanism remains controversial (reviewed in reference 359). It has been unclear how yeast WASp (Las17) activity is controlled. Las17 lacks a GBD and fails to bind yeast Rho GTPases (213), and there are no obvious yeast homologues to the proteins comprising the WAVE transinhibitory complex. Biochemically, purified full-length Las17 is active (i.e., not auto-inhibited) and constitutively 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, it seems likely that Las17 activity may be controlled by transinhibition, similar to WAVE1. In addition, Las17 has many other ligands 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 challenge will be to determine how these different factors temporally control Las17 activity at different stages of patch formation.

There also may be lessons to learn from studies in mammalian cells, where WIP binds stably to and inhibits N-WASp. This N-WASp-WIP complex is activated by Cdc42 specifically in the presence of Toca-1 (homologous to Bzz1), which binds to Cdc42 and N-WASp (151). All of the homologous proteins are expressed in yeast, making it tempting to draw direct parallels between these systems. However, Las17 and Bzz1 both lack Cdc42-binding domains. Further, bzz1Δ mutants show no obvious defects in cell growth, actin organization, or endocytosis (356). Thus, Las17 regulation by these factors may be distinct in yeast and/or require additional proteins to achieve effects similar to animal cell interactions of Cdc42-TOCA-WIP-WASp.

(ii) Pan1.

Pan1 is recruited to early patches along with Las17, well before the arrival of the Arp2/3 complex and detection of filamentous actin. Pan1 could be recruited to patches through its interactions with End3, Sla1, and/or clathrin adaptors (363, 364, 402). pan1Δ mutations are inviable, but mutation of the Pan1 acidic domain (pan1-101, which abolishes NPF activity) causes no obvious growth defects. This mutation does however show synthetic growth defects when combined with arp2 alleles (71, 87, 402). Thus, it appears that Pan1 performs essential functions required for endocytosis and cell growth distinct from its role as an NPF.

Pan1 NPF activity requires its A motif, which interacts directly 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 Arp2/3 complex to filament sides to stimulate nucleation and possibly promote formation of branched actin networks, such as those seen at patches (314, 423). It is unclear when in patch development Pan1 functions as an NPF. However, its NPF activity is strongly inhibited in vitro by Prk1 phosphorylation (371). These phosphorylation events could signal the observed loss of Pan1 from patches upon transition to rapid movements. Abp1 may also play a role in these events, as it arrives just before this transition and directly recruits Ark1/Prk1 kinases to patches (68, 102). Accordingly, abp1Δ cells show a delay in loss of Sla1 from patches as they transition to rapid movement (174). Presumably, Pan1 also remains on fast-moving patches in abp1Δ cells, since Sla1 and Pan1 form a complex that is disrupted by Ark1/Prk1 phosphorylation (427).

(iii) Myo3 and Myo5.

Just before the onset of rapid patch movement, Las17 and Pan1 are joined by the genetically redundant type I myosins Myo3 and Myo5 (172, 174). These proteins are comprised of an N-terminal motor domain, a lipid-binding TH1 domain, an F-actin-binding TH2 domain, an SH3 domain, and an Arp2/3 complex-binding A motif (Fig. (Fig.10).10). Myo3 and Myo5, like Las17, are required for actin patch assembly in the permeabilized cell assay (see “Las17/WASp” above), and myo3 myo5 double mutants are synthetic lethal with las17ΔWA mutants, suggesting that Myo3, Myo5, and Las17 are redundant as NPFs (98, 207). The NPF activities of Myo3 and Myo5 have not yet been characterized biochemically, but the TH2-SH3-A fragment of S. pombe myosin type I activates the Arp2/3 complex in vitro (210). This fragment binds with very weak affinity to F-actin and does not bind G-actin, leaving the myosin I NPF mechanism unclear. One model is that Myo3 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 Vrp1 enhances the NPF activity of the TH2-SH3-A fragment (352). In addition, it has been suggested that Myo3 and Myo5 function in a 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 electron microscopy in myo3 myo5 mutants (172), it has been proposed that Myo3 and Myo5 help promote the scission of endocytic vesicles from the membrane at the onset of rapid patch movement. Consistent with this possibility, fission yeast type I myosin can alter the composition and organization of membrane domains (361). How the motor activity of myosin contributes to function awaits determination, but motor activity is required for patch assembly in the permeabilized cell assay (207). Motor activity could produce a specific arrangement of actin filaments or, alternatively, transport a cargo such as the Arp2/3 complex to specific regions within a patch. Work on Acanthamoeba castellanii type I myosin (281) indicates a low duty ratio for type I myosins, suggesting that processive movement by Myo3/Myo5 requires multiple motors on a cargo.

(iv) Abp1.

Abp1 arrives at actin patches approximately when Myo3 and Myo5 do, just prior to the onset of rapid patch movement (173). However, once rapid movement commences, Myo3 and Myo5 are left behind at the cortex (172) while Abp1 remains associated with patches throughout internalization and transport on cables (159, 173). abp1Δ mutants fail to shed Sla1 and possibly other early patch components during the transition to fast patch movement, suggesting that Abp1 has a role in disassembly of the endocytic structures (174). Consistent with this model, Abp1 recruits Ark1 and Prk1 kinases to patches (68, 102); these kinases phosphorylate Pan1 to disrupt the Sla1-Pan1-End3 complex (427). This model also is consistent with Abp1's role in recruiting to patches Sjl1 (synaptojanin homologue), which is implicated in endocytic coat disassembly (357).

The biochemical properties of Abp1 provide a plausible explanation 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-terminal SH3 domain (Fig. (Fig.10).10). The NPF activity of Abp1 requires not only its A motifs but also its ADFH domain, through which it binds directly to F-actin with submicromolar affinity (127). These properties allow Abp1 to recruit the Arp2/3 complex to the sides of preexisting filaments, which leads to its modest NPF activity. The abilities of Abp1 to bind both F-actin and the Arp2/3 complex are required in vitro for Abp1 NPF activity and in vivo for its shared genetic functions with Sac6 (307). Similar to its mammalian relative cortactin (375, 399), Abp1 may displace Las17/WASp from the Arp2/3 complex and stabilize 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 with high affinity, yet has notably weak NPF activity compared to full-length Las17 and Pan1 (127). These properties are well-suited for Abp1 acting as a competitive antagonist, which is supported by Abp1 attenuation of Las17 NPF activity in vitro and by genetic interactions of abp1Δ with other NPF mutants (71). It is also consistent with the appearance of Abp1 at patches after other NPFs (173). Thus, sequential interactions of NPFs with the Arp2/3 complex may regulate the transition from slow to fast patch movement.

In addition to its NPF role, Abp1 recruits Srv2/CAP to patches by physically linking it to F-actin (23, 82, 214). As such, Abp1 may facilitate rapid turnover of F-actin, which is required for slow patch movements prior to the onset of rapid patch movement (173).

Coronin: spatial inhibitor of Arp2/3 complex activity.

A novel form of Arp2/3 complex regulation, distinct from NPF activity, is mediated by the yeast homologue of coronin Crn1, which localizes to actin patches (128, 142). Crn1 was isolated initially from crude extracts on microtubule affinity columns as a factor coeluting with actin (128). Recombinant Crn1 binds with high affinity to F-actin and cross-links F-actin to microtubules. F-actin binding is mediated by an amino-terminal WD repeat β-propeller structure, although the actin-binding surface has not been identified. This domain is followed by a 200-residue-long “unique” region of unknown structure that mediates microtubule binding and is not conserved in other species. At the C terminus is a coiled-coil domain that mediates Crn1 oligomerization and is required for F-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 β-propeller and coiled-coil domains are conserved in coronins from diverse species, as are the abilities to bind and cross-link F-actin (75).

The coiled-coil domain of Crn1 also binds directly to the Arp2/3 complex, through interactions with the ARPC2/p35 subunit. This interaction directly inhibits actin nucleation by the Arp2/3 complex (161), arising from direct Crn1-Arp2/3 interactions (NPF independent) that stabilize the Arp2/3 complex in its open (inactive) conformation (316). In electron micrographs of the Crn1-bound Arp2/3 complex, the globular β-propeller domain rests ∼20 Å from the ARPC2/p35 subunit (Fig. (Fig.9A).9A). The coiled-coil and unique regions mediating this interaction are not visible, presumably because they lack substantial mass and/or tertiary structure. Since the mammalian coronin Cor1 also coimmunoprecipitates with the Arp2/3 complex, this regulatory function of coronin may be widely conserved (50).

What is the cellular function of Arp2/3 complex inhibition by Crn1? While the coiled-coil domain of Crn1 mediates inhibition, the presence of preformed actin filaments relieves inhibition and thereby permits Arp2/3 nucleation activity (161). Thus, Crn1 appears to spatially restrict Arp2/3 activity to the sides of preexisting filaments by suppressing de novo nucleation in the absence of filaments. As such, Crn1 may promote formation of branched filament networks, a model that can be tested by characterizing Crn1 effects on filament branching by the Arp2/3 complex.

Crn1 localization to cortical patches requires both its β-propeller and coiled-coil domains, suggesting requirements for both F-actin binding and coiled-coil-mediated functions (oligomerization and/or Arp2/3 complex binding). Although the timing of its arrival to patches has not been reported, it may be similar to that of other F-actin-binding proteins, Abp1, Sac6, and Cap1/Cap2, arriving late and persisting on patches through rapid movements. Surprisingly for a factor with such potent inhibitory activity on the Arp2/3 complex, deletion of CRN1 does not result in detectable growth or morphological defects (128). However, a physiological role for Crn1 in regulating the Arp2/3 complex is supported by synthetic defects between crn1Δ and arp2-21 mutations (161). Further, crn1Δ has synthetic defects in cell growth and actin organization with cof1-22 and act1-159 (mutations that slow actin turnover), suggesting a role 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 Arp2/3 complex activity contribute to patch maturation and endocytosis? Mechanistically, how does the presence of F-actin override Crn1 inhibition of the Arp2/3 complex? Does Crn1 regulate actin turnover in vivo with cofilin? Does Crn1 interact with microtubules in vivo, and how might this be coordinated with its actin functions?

Open questions about the Arp2/3 complex.

In addition to direct inhibition of the Arp2/3 complex by Crn1, binding of certain proteins (e.g., tropomyosin) to actin filaments prohibits Arp2/3 complex association and thus indirectly inhibits the Arp2/3 complex (37). This may explain why the Arp2/3 complex is not detected on actin cables, which are decorated by Tpm1 and Tpm2. It also raises an important mechanistic question: how do cables and not patches selectively become decorated with Tpm1 and Tpm2? Another factor that may regulate Arp2/3 complex function is Cmd1 (calmodulin), which interacts both genetically and biochemically with the ARPC2/p35 subunit (330). There are likely many other cellular factors that must be coordinated with NPFs and Crn1 to control Arp2/3 complex function.

There are also many open questions regarding Arp2/3 complex cellular function. First, what is the specific role of Arp2/3 complex-based actin polymerization in patch maturation and endocytosis? The Arp2/3 complex is required for actin assembly, and actin assembly is required for endocytosis, but what function does this provide? Actin polymerization could provide force generation to drive vesicle scission; alternatively, it could provide a dynamic scaffold to organize endocytic factors that drive membrane invagination and scission (93). There are also non-patch/endocytosis functions of the Arp2/3 complex to consider. For instance, the involvement of the Arp2/3 complex in cable and cytokinetic ring assembly has not been ruled out. Although studies on arp2, arp3, and arc35 mutants have not indicated a role in cable assembly (99, 407, 408), given the physical links between patches and cables (159, 177) and multiple reports suggesting interactions between the Arp2/3 complex and formins (69, 152), this question deserves further inquiry. Further, while one study in budding yeast suggests that the actin ring is assembled by formins and not the Arp2/3 complex (369), a separate study in fission yeast reports a role for the Arp2/3 complex in ring assembly (288). Thus, additional investigations are needed to resolve this potential Arp2/3 complex function at cytokinetic rings. Finally, separate from assembly of the three major actin structures—patches, cables, and rings—Arp2/3 complex function contributes to organelle inheritance, nuclear export, and mitotic spindle orientation (41, 330, 331, 416). Understanding the mechanistic basis of these functions could broaden the range of action for the Arp2/3 complex.

Assembly of Actin Cables

Formins Bni1 and Bnr1.

Actin cables are essential for polarized cell growth, but until recently the mechanism of actin cable assembly was unknown. Early cell biology studies suggested a role for the two budding yeast formins Bni1 and Bnr1 in cable formation (97, 164, 176, 182, 284), but it was hypothesized they had indirect roles in this process, serving as molecular scaffolds. Further, technical obstacles in purifying these large proteins (∼200 kDa) delayed characterizing their effects on actin in vitro. Similar to all formins of the Diaphanous-related formin (DRF) family, Bni1 and Bnr1 are comprised of well-defined domains that differentially contribute to their mechanism and regulation (Fig. 11A). These include the formin homology 2 (FH2) domain, FH1 domain, GBD, Diaphanous inhibitory domain (DID), and Diaphanous autoregulatory domain (DAD) (100, 389). A critical breakthrough in understanding the role of formins in regulating the actin cytoskeleton came when two studies showed that the FH2 domain of Bni1 is critical for cable formation (99, 325). Biochemical studies closely followed, demonstrating that purified FH2-containing fragments of Bni1 directly nucleate actin assembly (303, 326). Similar actin-nucleating activities have since been reported for formins from a wide variety of animals, fungi, and plants, making it clear that actin assembly is a conserved function of formins (149).

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 ...

Budding yeast expresses two formins, Bni1 and Bnr1 (164), which partially overlap in their localization to the bud tip and bud neck (Fig. (Fig.5A).5A). Bni1 and Bnr1 also have distinct and overlapping functions in the assembly of cables and the actomyosin ring (99, 304, 325, 369). A comparison of their biochemical activities reveals differences: Bnr1 specifically bundles actin filaments and has 10- to 15-fold more potent actin assembly than that of Bni1 in vitro (251). Bni1 and Bnr1 also appear to be regulated by distinct mechanisms (see below). How these differences in activity and regulation tailor Bni1 and Bnr1 to their respective cellular functions remains to be determined.

What is the formin mechanism of actin assembly? Whereas the Arp2/3 complex bypasses kinetic barriers to actin assembly by mimicking the barbed end of an actin filament, formins use a different mechanism. Kinetic studies suggest that the FH2 domain promotes filament assembly by stabilizing polymerization intermediates, actin dimers and/or trimers, which are otherwise extremely short-lived species (300). This is facilitated by formins having very high affinity for F-actin. Once nucleated, formins remain bound to the rapidly growing barbed end of the filament while guiding insertional addition of actin subunits. This activity is referred to as processive capping. The first indications of this activity came from the demonstration that Bni1 FH2 localizes to filament barbed ends by immunoelectron microscopy (303) and does not completely inhibit barbed end growth (300, 303). Two subsequent studies showed that Bni1 FH2 protects elongating barbed ends of filaments from the inhibitory effects of conventional capping proteins (253, 429). This process, which involves a strong interaction between FH2 and filament barbed ends (Kd ≤ 20 nM) (253, 300, 303), has now been visualized in real time using TIRF microscopy of individual FH2-capped actin filaments (197) and drives rapid barbed end filament growth at various rates (194, 197, 300, 303, 429).

While a thorough mechanistic understanding of processive capping is still far off, critical insights have been gained by recent structural analyses. First, the FH2 dimerizes, and point mutations disrupting dimerization abolish actin assembly (253). This suggested that a two-headed actin-binding structure was required, opening the possibility of alternating actin-binding contacts to allow insertional growth and persistent association with the filament barbed end. Shortly thereafter, the crystal structure of the Bni1 FH2 domain was solved, which revealed a flexibly tethered dimer in which each FH2 molecule has two actin-binding sites (Fig. 11B) (414). Sites of contact between FH2 and actin have since been defined at an atomic level by a cocrystal structure of the Bni1 FH2 domain bound to tetramethylrhodamine-labeled actin (282) (Fig. 11C).

Based on these findings, several different models for processive capping have been proposed. First, the stair-stepping model envisions two flexibly tethered actin-binding heads alternating interactions with the growing barbed end of the filament (414). Anticooperative conformational changes in FH2 drive alternating contacts to allow subunit addition (414). A caveat to this mechanism, however, is that it requires rapid rotation of the FH2 dimer to account for a helical twist of the actin filament, and experimental evidence suggests that such a rotation may not occur (197). To resolve this paradox, a second model suggests that the unidirectional twist induced by multiple stair steps may be relieved by a periodic “backward” step, in which the FH2 twists in the opposite direction as the filament (344). A third model proposes that the FH2 dimer may act like a shaft in a bearing, encircling the growing actin filament (197). However, the molecular contacts that maintain persistent FH2 association with the growing barbed end in this model are not clear. Finally, a fourth model suggests that the FH2 may adopt an end-side-binding mechanism, whereby it associates with the side of barbed ends to allow new subunit addition while prohibiting binding of capping proteins (137). Experimental evidence has not yet ruled out any of these mechanisms and, in fact, they may be more similar mechanistically than they are distinct. Resolving the mechanism may require the use of high-resolution electron microscopy techniques to image FH2 dimers bound to the barbed ends of filaments. Similar techniques have been used to define conformational changes in the Arp2/3 complex upon WASp activation and at filament branches (91, 316) and to image protein complexes at microtubule ends (422).

While recent efforts have focused on defining the formin mechanism of actin assembly, many critical questions remain open surrounding formin regulation. Like WASp and the Arp2/3 complex, the potent activities of formins are likely subject to tight spatial and temporal regulation. Below, we discuss what is known about the regulation of Bni1 and Bnr1 by their known ligands, which include Rho GTPases, profilin, Bud6/Aip3, Tef1 (eEF1α homologue), and Spa2.

Profilin-FH1 interactions: a throttle for filament elongation.

Profilin is an abundant actin monomer-binding protein found in all eukaryotic cells examined. Further, profilin-bound actin monomers are considered the primary physiological substrate available for actin assembly in vivo. Profilin has two separate binding surfaces, one that binds actin monomers and another that binds polyproline motifs, including those found in FH1 domains (225, 410). Indeed, early studies identified profilin as a ligand of Bni1 and Bnr1 (97, 164). Profilin binding blocks actin monomer addition to the pointed ends of filaments but freely allows addition to barbed ends (297), consistent with the profilin-actin cocrystal structure (Fig. (Fig.8)8) (336). These properties of profilin are fundamental to how it contributes to the formin mechanism of actin assembly.

Importantly, the Arp2/3 complex nucleates actin filaments with free barbed ends and, thus, Arp2/3-nucleated filaments elongate at similar rates in the presence and absence of profilin (315). In contrast, formins remain tightly associated with the barbed ends of filaments they nucleate and sterically inhibit capping protein association, raising an important mechanistic question: can profilin-bound actin monomers be added efficiently to the barbed ends of formin-capped filaments? The first insights came from Sagot et al. (326), who found that Bni1 FH1-FH2 assembles actin filaments equally well from a pool of free actin monomers or from profilin-bound actin monomers. Further, FH1-profilin binding is required for formins to assemble profilin-bound actin monomers (300, 326). Thus, FH2 alone promotes filament assembly from free actin monomers, but FH1 is required for filament assembly from profilin-bound actin monomers. This emphasizes the importance of the FH1 domain for formin function in cells, where profilin-bound actin monomers are the primary substrate available for filament assembly. As such, FH1 is required for formin-mediated actin assembly in vivo, which is supported by many studies (99, 148, 196, 253, 303, 325, 326, 392).

More-recent biochemical studies have revealed exciting effects on actin assembly resulting from profilin-FH1 interactions with profound implications for formin function in vivo. By studying an isoform of mammalian formin mDia1 with extended polyproline tracts in its FH1, Romero and coworkers demonstrated a formin/profilin-dependent increase in rate of barbed end elongation—many times that of the diffusion-limited rate of free barbed end growth (321). Prior to this work, it was a widely held view that actin polymerization could not exceed diffusion-limited rates of assembly (∼11 subunits per second per micromolar). Thus, in a solution of 10 μM G-actin, filaments would elongate at ∼110 subunits per second. The work of Romero et al. showed that filaments capped by formins can elongate at least fivefold faster specifically when profilin is present. A more recent study now has used TIRF microscopy to elegantly demonstrate these effects in real time for individual actin filaments capped by a variety of formins, including Bni1 (194). Further, they show that the magnitude of the increase in filament elongation rate correlates with the number of profilin-binding sites in FH1. The largest increase measured (∼5-fold) occurs on filaments capped by mDia1 isoforms 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 elongation at free barbed ends). Bnr1 has not yet been examined, but it has two predicted profilin-binding sites, suggesting it likely increases the rate of elongation by no more than twofold.

These results have important implications for formin function in vivo. The yeast actin cable flow rate is ∼0.3 μm/s (419). If cable flow is driven primarily by actin polymerization, as suggested, this corresponds to addition of ∼100 subunits/s at filament ends, requiring a cellular concentration of ∼9 μM G-actin. However, the total concentration of actin in yeast cells is estimated to be 12 μM (A. Goodman and B. Goode, unpublished data), the majority of which is thought to be in filamentous form (178). Thus, the G-actin concentration in yeast cytosol should be well below 5 μM. These discrepancies can begin to be reconciled by considering the formin/profilin-dependent increase in elongation rates. Further, this may explain why measured rates of actin assembly at patches (0.05 to 0.1 μm/s) are markedly lower than cable flow rates (173, 174).

Although it is now clear that the FH1 domain serves as a throttle controlling the speed of elongation at formin-capped filaments, the mechanism remains to be determined. One possibility is that profilin presents monomers in an orientation optimal for their addition to barbed ends. It is thought that only about 2% of random collisions between an actin monomer and the free barbed end of a filament result in effective docking and addition (83). FH1-profilin-G-actin interactions may increase the efficiency of this step. A second possibility, not mutually exclusive from the first, is that profilin-FH1 interactions may increase the concentration of G-actin near the barbed end of the filament to increase the rate of elongation. The stage is set for testing these models. Another critical aspect of the mechanism that should be tested is whether the FH1 directly transfers profilin-bound monomers to the barbed end, 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-hybrid screen for actin-interacting proteins (13) and a genetic screen for mutants defective in the bipolar budding pattern (426). Bud6 localization in cells is similar to that of Bni1 (14, 284), and bud6Δ cells show defects in formin-mediated actin cable assembly (99, 325). Further, in two-hybrid assays Bud6 has been shown to interact with Bni1 (97) and Bnr1 (182). Subsequent biochemical work demonstrated that the C-terminal half of Bud6 (i) binds specifically to actin monomers but not filaments, (ii) stimulates nucleotide exchange on G-actin, and (iii) enhances Bni1-mediated actin assembly in vitro additively with profilin (253). Consistent with these effects, bud6Δ was found to be synthetic lethal with pfy1-4, suggesting that Bud6 and profilin have related and/or complementary cellular functions. Subsequently, the Bud6-binding site was mapped to a short sequence in the C terminus of Bni1, overlapping with its DAD (251), raising the possibility that Bud6 may participate in activating Bni1 from an inhibited state (see below). Although Bud6 interactions with both Bni1 and Bnr1 were detected in the two-hybrid studies above, biochemical assays could detect interactions of Bud6 specifically with Bni1 and not Bnr1. Further, mutational analysis revealed that key residues in Bni1 required for Bud6 binding are not conserved in Bnr1. Actin assembly assays confirmed this differential interaction; Bud6 stimulates Bni1 activity by ∼2- to 3-fold but has no effect on Bnr1 activity. Further mechanistic studies will be required to determine precisely how Bud6 enhances Bni1-mediated actin assembly, which in principle could arise from enhanced nucleation and/or elongation.

Multiple lines of genetic evidence indicate additional cellular functions for Bud6 beyond Bni1 stimulation. For example, bud6Δ cells have more severe defects in actin organization than bni1Δ cells, suggesting that Bud6 may contribute (directly or indirectly) to the regulation of Bnr1 and/or other actin organizing factors. Further, Bud6 has roles in capturing astral microtubules to regulate mitotic spindle orientation (160, 338, 339) and in organization of the endoplasmic reticulum at the bud neck to compartmentalize mother and daughter cells (226).

Rho GTPases.

The localized assembly of yeast actin cables and cytokinetic rings by Bni1 and Bnr1 necessitates tight spatial and temporal control of their activities. Thus, one critical question is how Bni1 and Bnr1 actin assembly activities are up-regulated and down-regulated. Work on mammalian formins mDia1 and mDia2 has shown that they are auto-inhibited by interactions between their amino-terminal DID motif and their carboxyl-terminal DAD region (8, 392) (Fig. 11A), and these interactions abolish actin nucleation activity of the FH2 (212). Many formins also contain an amino-terminal GBD adjacent to the DID (211, 322). Direct binding of Rho GTPases to the GBD is proposed to release formins from an auto-inhibited state (8, 392). However, inhibition of mDia1 FH2 activity by DID is only minimally relieved by addition of RhoA (211, 212), suggesting other and/or more mechanisms may be required for formin activation.

The overexpression of N- and C-terminally truncated Bni1 and Bnr1 constructs produces excess actin assembly in vivo (97, 99, 304, 325), which has led to the suggestion that Bni1 and Bnr1 may be auto-inhibited like mDia1 and mDia2. This is consistent with conservation in Bni1 (and to a lesser degree Bnr1) of the key functional residues in DAD (8). However, it should be noted that the critical DAD-binding residues in DID are not conserved in Bni1 (203, 267) and that direct interactions between N and C termini have not been reported for either Bni1 or Bnr1. This leaves open the question of whether Bni1 and/or Bnr1 is auto-inhibited, which should be addressed through biochemical analyses of full-length proteins.

Regardless of whether Bni1 and Bnr1 are down-regulated by auto-inhibition or alternative mechanisms (e.g., transinhibition similar to that of Las17), Rho GTPases play key roles in yeast formin regulation in vivo. Six different Rho family GTPases are expressed in budding yeast: Rho1, Rho2, Rho3, Rho4, Rho5, and Cdc42. Bni1 interacts with Cdc42, Rho1, and possibly Rho3 and Rho4 (97, 190), while Bnr1 binds Rho4 (164). Bni1 localization depends on Cdc42 (284, 304). Further, Dong and coworkers have addressed the functional importance of formin-Rho interactions by testing the ability of N-terminally truncated (activated) Bni1 and Bnr1 constructs to suppress defects caused by rho mutants at different stages of the cell cycle (79). Their work implicates Cdc42 in the control of formin-mediated actin assembly at bud emergence, Rho3 and Rho4 during bud growth, and Rho1 during the response to cell stress. Another study implicates Rho1 in regulating Bni1-mediated actin assembly at the cytokinetic ring (369). These studies have made an important contribution by providing working functional pathways for formin regulation by Rho GTPases, but they leave open the possibility that some of the observed effects are not direct. Thus, biochemical evidence for the effects of Rho GTPases on formin activity is needed to clarify the functional roles of these interactions.

Other Bni1 and Bnr1 ligands.

A number of other cellular factors interact with Bni1 and/or Bnr1, for which the functional implications of the interactions are poorly understood. Tef1 (eEF1α homologue) is a highly abundant translation elongation factor that interacts with Bni1 FH2 (residues 1328 to 1513) (374), binds to and bundles F-actin in vitro (76, 262), and contributes to actin cable organization (132). Binding of a Bni1 fragment to Tef1 disrupts Tef1 bundling of F-actin (374), but the effects of Tef1 on Bni1 activity have not been investigated. Interestingly, overexpression of TEF1 induces formation of supernumerary actin cables and enlarged depolarized cells (261), similar to the effects of overexpressing dominant formin constructs. As Tef1 is one of the most abundant proteins in yeast cells (55) and may link functions of the actin cytoskeleton and translational control, it will be of great interest to define the effects of the Bni1-Tef1 interaction.

Additional factors that may affect formin function include Hof1, Spa2, and Smy1. Hof1, an SH3 domain-containing protein of the PCH family (216), interacts with Bnr1 FH1 by the two-hybrid assay and localizes to the bud neck similarly to Bnr1 (176). Genetic data also suggest that Hof1 may regulate Bnr1 function at the bud neck (215, 378). Hof1 is proposed to mediate septum formation to facilitate cytokinesis, rather than actomyosin ring formation (33), and thus Hof1 could regulate Bnr1-mediated cable assembly to promote septation. Another formin ligand, Spa2, appears to interact with Bni1, as Bnr1 lacks the Spa2-binding domain (113) (Fig. 11A). Spa2 is required for proper localization of Bni1 (284), but its potential effects on Bni1 activity have not been tested. The kinesin-related protein Smy1 interacts by two-hybrid assay with Bnr1 FH2 (182), but no functional role for this interaction has yet been reported. Intriguingly, Smy1 also interacts with type V myosin Myo2 (28), providing a possible link between cable assembly by Bnr1 and transport on cables.

Organization and Stabilization of Actin Filaments

Bundling proteins.

The distinct functions of cellular actin arrays require organization of actin filaments into specialized architectures (e.g., patches, cables, and rings). This is regulated by proteins that bundle, cross-link, and/or stabilize filaments into specialized configurations. Below, we discuss the known mechanisms and cellular functions of yeast actin-binding proteins 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), each comprised of a pair of calponin homology (CH) domains (Fig. (Fig.12).12). True to its initial codiscovery by genetic (4) and biochemical (84) approaches, a deeper understanding of the Sac6-actin interaction has developed through the combination of genetic and biochemical studies. Initially, SAC6 was identified as an extragenic suppressor of temperature-sensitive act1 alleles, in which point mutations in Sac6 suppressed the growth defects caused by point mutations in actin (4, 5). Independently, Sac6 was isolated on F-actin affinity columns and shown to localize in vivo to patches and cables (84). Then, it was demonstrated that Sac6 bundles F-actin in vitro and is required for proper actin organization in vivo (3). Subsequent genetic and biochemical analyses demonstrated that Sac6 contacts residues on the surface of actin subdomains 1 and 2 exposed on the “edge” of the actin filament (46, 154). Further, some of the defects observed in sac6Δ cells (e.g., synthetic lethality with abp1Δ and sla2Δ) are mimicked by mutations at the Sac6 contacts in actin subdomains 1 and 2 (153, 329). To date, the cognate actin-binding surfaces on Sac6 have not been identified, despite the availability of Sac6/fimbrin crystal structures to guide mutational analyses (123, 189).

FIG. 12.
Actin filament-bundling proteins. Schematics of each protein are drawn to scale. Abbreviations: CC, coiled-coil; CLR, calponin-like repeat; COOH, carboxyl-terminal domain. Known actin-binding domains are underlined. Yeast IQGAP (Iqg1) is included because ...

All actin-bundling proteins cross-link filaments by one of two mechanisms: (i) having multiple ABDs or (ii) oligomerization. Sac6 bundles filaments by having two separate ABDs. High-resolution EM studies on Sac6-decorated filaments agree with previous genetic work defining the Sac6-actin interface and provide a model for the cross-link (135, 136, 386). Until recently, it was thought that Sac6 was not regulated and that its two actin-binding heads were available to interact with filaments. However, recent crystal structures of A. thaliana and S. pombe Sac6/fimbrin reveal a “closed” conformation, in which the two ABDs fold in on each other, preventing exposure of residues hypothesized to mediate actin binding (189). This suggests that an induced structural shift may be required for actin binding to occur. Consistent with this possibility, distinct conformations for Sac6 have been observed by EM and crystallographic studies, but how such rearrangements might be regulated is not yet clear. One possibility is that the EF-hand motif in Sac6/fimbrin (Fig. (Fig.12)12) allows regulation by calcium and/or other signaling molecules.

While key features of the Sac6-actin interaction now are understood, the cellular function(s) of Sac6 remains unclear. For instance, Sac6 function in endocytosis has not been resolved. Actin patches in sac6Δ cells are impaired in moving inward from the cell cortex (174), suggesting that Sac6 may contribute to actin-based force generation, but the mechanism of this contribution is unknown. Further, how does a bundling protein associate with a network of branched filaments such as an actin patch? Perhaps Sac6 bundles filaments at specific stages in patch maturation, either prior to or subsequent to the stage characterized by EM where bundles are not obvious (314, 423). Additionally, Sac6 bundling may be used to link patches and cables during fast patch movement on cables (159). Another possibility that should be considered is that the abilities of Sac6 to bundle and stabilize filaments may be separate. Thus, Sac6 could function primarily to stabilize rather than bundle filaments in patches. Consistent with this model, partially purified actin patches isolated from sac6Δ cells are less stable than those isolated from wild-type cells (423). To resolve these issues, mutants that specifically impair each of the ABDs of Sac6 should be generated, and actin patch ultrastructure in sac6Δ mutants should be analyzed by EM to determine whether there are visible defects in filament organization.

The precise function of Sac6 in actin cables is also unknown. sac6Δ cells show diminished cable staining and rounded cell morphologies, indicative of polarity defects (3). Thus, Sac6 may bundle and/or stabilize the actin filaments in cables. However, this has not been tested directly, for instance, by measuring rates of cable turnover in sac6Δ cells. Finally, a functional role for Sac6 at the actomyosin ring has not been investigated. S. pombe fimbrin localizes to the medial ring and functions during cytokinesis (412), raising the possibility that Sac6 functions similarly.

(ii) Scp1.

Scp1 (a calponin/transgelin homologue) is the only other component of patches or cables (other than Sac6) that contains a CH domain; in addition, the bud neck component Iqg1 contains a CH domain (see below). Like Sac6, Scp1 localizes to actin patches and bundles and stabilizes actin filaments in vitro (129, 405). Deletion of SCP1 exacerbates defects caused by deletion of SAC6, further suggesting their functional relationship. However, biochemical dissection of Scp1 activity reveals clear mechanistic differences between these two bundlers. First, Sac6 bundles filaments through the use of two ABDs, each comprised of a pair of CH domains. However, Scp1 bundles filaments through C-terminal sequences distinct from its CH domain (Fig. (Fig.12).12). This relatively small C-terminal region of Scp1 is both required and sufficient for bundling but does not appear to oligomerize, leaving its bundling mechanism uncertain. One possibility is that this fragment harbors two separate ABDs. Alternatively, actin binding may alter the conformation of Scp1 to promote oligomerization (similar to IQGAP [see below]). Scp1-induced actin filament bundles also are qualitatively distinct from Sac6-induced bundles, consistent with their apparently divergent actin interactions.

Scp1 has not been visualized on actin cables or rings, but its relatively low abundance compared to Sac6 may preclude visualization on those structures. Indeed, even the more abundant Sac6 shows only faint localization on cables (84). Thus, it is an open possibility that Scp1 and Sac6 function together to regulate cables. Scp1 also is linked to formation of reactive oxygen species regulating yeast cell death (130), but it is not yet clear whether this function requires Scp1 interactions with actin. Key to understanding Scp1 cellular function will be determining the respective roles of its N-terminal CH domain and C-terminal actin-bundling domain. It is possible that the CH domain functions as a “locator” domain to facilitate activity of the C terminus. Alternatively, the CH domain may mediate interactions with other cellular factors. Thus, future work on Scp1 should (i) define which domains of Scp1 are required for its localization and (ii) identify new Scp1 ligands.

(iii) Iqg1/Cyk1.

A third CH domain-containing protein in yeast is Iqg1 (also called Cyk1), an IQGAP homologue. Most IQGAP family members have a CH domain at their N terminus, calmodulin-binding IQ motifs, and a C-terminal Ras-GAP domain (RGD). The RGD mediates interactions with Cdc42 but lacks the GAP activity originally proposed (Fig. (Fig.12)12) (228). Cdc42 and Iqg1 two-hybrid interactions have been reported (279), but interactions could not be detected biochemically between Iqg1 and any of the yeast Rho GTPases (96, 341), leaving open whether Iqg1 is regulated in this manner. The IQG1 gene is essential for yeast cell growth, and Iqg1 localizes to the bud neck in a manner dependent on its IQ motifs and their interactions with Mlc1 (96, 217, 342). The terminal phenotype of iqg1Δ cells suggests that it plays a critical role in cytokinesis.

Iqg1 binds to F-actin through its N terminus, which contains the CH domain (96, 341). Filament bundling by Iqg1 has not been tested. However, mammalian IQGAPs bundle filaments, and this activity is regulated by their interactions with Cdc42 and calmodulin. Cdc42-GTP enhances bundling, possibly by promoting IQGAP oligomerization (114), whereas calmodulin inhibits bundling (26). One implication from these studies is that IQGAP likely bundles via oligomerization, rather than utilizing multiple ABDs on a single polypeptide chain. Since Iqg1 interacts with calmodulin and Mlc1 (341, 342), and even possibly Cdc42, it may be regulated in a similar manner, but this awaits future tests.

What specific cellular function does Iqg1 perform to promote cytokinesis? Truncation analyses show that the CH domain contributes to assembly of the actomyosin ring, that the IQ motifs are required for Iqg1 recruitment to the bud neck (via interactions with Mlc1), and that the RGD facilitates contraction of the actomyosin ring (341). This puts Iqg1 at the center of a fascinating network of interactions driving cell division. However, the mechanisms by which Iqg1 facilitates actin ring formation and subsequent ring contraction remain unclear. Does the CH domain interact directly with actin filaments to promote ring formation, or are there other ligands involved? What molecular interaction(s) of the Rho-binding domain promotes ring contraction?

(iv) Crn1.

Here, we discuss the actin-binding and -bundling activities of Crn1 (yeast coronin); its role in regulating the Arp2/3 complex is described above. The N terminus of Crn1 forms a β-propeller domain that mediates a high-affinity interaction with F-actin; the C terminus contains a coiled-coil domain (Fig. (Fig.12).12). Purified Crn1 oligomerizes via its coiled-coil domain and cross-links filaments into either loose arrays of overlapping short filaments or smooth bundles of longer filaments (128). Overexpression of Crn1 in yeast causes arrest of cell growth, which can be suppressed by truncating either the ABD or the coiled-coil domain. Crn1 localization to actin patches also depends on both domains (161). Thus, Crn1 localization and function may depend on its ability to bundle F-actin. Alternatively, this may reflect a requirement for Crn1 interactions with the Arp2/3 complex, also mediated by the coiled-coil domain (161). Since there appear to be separate pools of Crn1 in cells—about 50% of which is associated with the Arp2/3 complex (161)—Crn1 may be performing multiple functions that involve its coiled-coil domain.

Deletion of CRN1 causes no obvious defects in cell growth, morphology, or actin organization, suggesting its functions are redundant with other cellular factors. However, Crn1 shows no genetic interactions with other bundling proteins (Sac6, Scp1, and Abp140). Instead, crn1Δ displays synthetic defects in cell growth and actin organization with act1-159 and cof1-22. This raises the intriguing possibility that Crn1 may promote actin turnover rather than stabilization (128). These and other questions regarding Crn1 cellular function need to be resolved.

(v) Abp140.

Abp140 is one of only two actin-binding proteins identified in yeast with no obvious mammalian counterpart, the other being Bud6 (Table (Table1).1). Abp140 was first purified from crude yeast cell extracts as a 140-kDa protein with actin-bundling activity (17). However, the ABP140 open reading frame predicts a protein with a calculated mass of 71 kDa. This suggests either that Abp140 has abnormal properties that cause retarded migration on sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels or that it oligomerizes in a manner resistant to the denaturing conditions of sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels. The migration of Abp140 on gel filtration columns is consistent with oligomerization (17). Thus, while the ABDs of Abp140 have not yet been identified, this protein appears to multimerize, which offers a potential mechanism for bundling. Abp140 localizes to both actin patches and cables. However, abp140Δ cells display no defects in cell growth, morphology, or actin organization. Genetic interactions of Abp140 with other actin-binding factors have not been tested, leaving its cellular function unclear.

Since its initial characterization, Abp140 has been used as a GFP-fused marker for studies monitoring actin cable dynamics (104, 159, 419). As multiple bundling proteins colocalize with patches and cables (e.g., Abp140 and Sac6 and possibly Scp1), an important mechanistic question is whether these factors compete for and/or cooperate in binding F-actin and help connect patches and cables. In addition, cable architecture likely is shaped by the activities of multiple bundling factors working in concert.

(vi) Tef1 and Tef2.

Tef1 and Tef2 are the yeast homologues of eukaryotic translation elongation factor 1α (EF1α), and they bear 81% sequence identity to their human counterpart (374). Tef1 and Tef2 are identical proteins, encoded by genetically redundant genes, that together are essential for cell viability (333). In addition to their well-documented roles in protein translation, all EF1α family members examined (from yeast to humans) bundle F-actin in vitro (67). In addition, Xenopus laevis and human EF1α proteins sever microtubules (347). Thus, Tef1 has the potential to coordinate protein translation with functions of both the actin and microtubule cytoskeletons.

Filament bundling was first demonstrated for Dictyostelium discoideum EF1α (418) and more recently for Tef1 (261). The bundling mechanism of Tef1 likely involves two separate ABDs, which have been mapped to two distinct 50-residue stretches, located at the N and C termini of Dictyostelium EF1α (218). EF1α bundles actin filaments into a unique architecture that excludes other cross-linking proteins from binding (283). Point mutations that disrupt the ability of Tef1 to bundle filaments, but leave its translation function unaffected, cause defects in actin cable organization in vivo (132). However, it remains unknown whether Tef1 localizes to cables or other F-actin structures, and thus it is unclear how it contributes to cable organization. As discussed above, Tef1 interacts with the actin-nucleating FH2 domain of Bni1, and this interaction disrupts F-actin bundling by Tef1 (374). However, it remains to be determined whether this interaction affects the actin cable assembly-promoting activity of Bni1. Another reported ligand of Tef1 is the actin monomer-binding protein Srv2/CAP (see below) (417), but the in vitro and in vivo consequences of this interaction remain unknown.

Finally, it will be interesting to learn how the functions of Tef1 in regulating the actin cytoskeleton are coordinated with its classical role in translation. The role of EF1α in translation requires GTP binding and hydrolysis (168, 312). F-actin binding inhibits the abilities of EF1α to bind GTP but promotes GTP hydrolysis (90). Thus, actin binding may drive EF1α into the GDP-bound form, inactive for translation.

Nonbundling Actin Stabilizers

(i) Tropomyosins Tpm1 and Tpm2.

Budding yeast express two tropomyosin isoforms, TPM1 and TPM2, which function primarily to stabilize the filaments in actin cables and the actomyosin ring. Intriguingly, they are the only proteins known to localize exclusively to cables, i.e., not also label patches. Tpm1 and Tpm2 have genetically overlapping yet possibly distinct cellular functions. TPM1 expression levels are ∼6 times higher than TPM2 levels in exponentially growing cells (81), which correlates with the relative severity of their phenotypes. Whereas tpm1Δ cells lack readily detectable cables, have cell morphology defects, and grow slowly (219), tpm2Δ cells show no obvious defects in growth, morphology, or actin organization (81). Further, overexpression of TPM2 rescues most of the defects caused by tpm1Δ (D. Pruyne and A. Bretscher, personal communication). The observation that tpm1Δ and tpm2Δ mutations are synthetic lethal is consistent with genetic redundancy in their functions but also leaves open the possibility that they perform some specific roles, as suggested by their biochemical differences (below).

Tpm1 and Tpm2 display many of the defining biochemical properties of tropomyosin family members, including dimerization, divalent cation-dependent F-actin binding, and resistance to denaturation at high temperatures (90°C) (61, 81, 220). However, Tpm1 and Tpm2 have different actin-binding stoichiometries (1 Tpm1 dimer per 4.6 actin subunits; 1 Tpm2 dimer per 3.6 subunits). Further, Tpm2 can displace Tpm1 from actin filaments, but Tpm1 fails to displace Tpm2 from filaments. Additionally, low levels of Tpm1 may actually enhance Tpm2 binding (81). This suggests that Tpm2 may have stronger affinity for F-actin and that its actin-binding interaction is qualitatively distinct from that of Tpm1. A future challenge is to determine how these biochemical differences translate into different cellular functions. This will likely require careful analysis of cable dynamics in tpm1 and tpm2 mutants and identification of differential genetic interactions for these mutants.

As discussed earlier, the generation of a fast-acting temperature-sensitive strain, tpm1-2 tpm2Δ, greatly facilitated the discovery that tropomyosin is required for cable stability (306). Tpm1 and Tpm2 may stabilize cables by prohibiting depolymerization/turnover factors, such as cofilin, from associating with filaments. Consistent with this view, filament binding by tropomyosin and cofilin are mutually exclusive in vitro and in Caenor habditis elegans muscle cells (30, 278). This is also consistent with the observation that yeast patches and cables are decorated differentially by cofilin (patches) and tropomyosin (cables). As actin cables are highly dynamic structures that undergo rapid filament turnover (19, 419), it will be important to determine whether tropomyosin-actin interactions are regulated. It seems likely that maintaining rapid turnover of cables would require active mechanisms to displace these filament stabilizers. Indeed, such mechanisms have begun to emerge; Tpm1 (and likely Tpm2) is acetylated directly by the Mdm20-containing NatB acetyltransferase complex, and this modification promotes Tpm1 binding to F-actin and cable stability in vivo (146, 292, 350, 351). This and other modifications, as well as competitive inhibitor mechanisms, may be required to maintain the delicate balance between cable stability and rapid turnover.

(ii) Capping protein.

All of the filament-stabilizing factors discussed above (e.g., Sac6, Scp1, and Tpm1/Tpm2) decorate sides of filaments and reduce rates of subunit dissociation as filaments shorten from their barbed and/or pointed ends. However, factors that directly target filament ends can also have tremendous stabilizing effects. It is difficult to find a more classic example of this than the CapZ/conventional capping protein family, of which yeast capping protein (Cap1/Cap2) is a member. These heterodimeric proteins tightly associate with and cap the barbed ends of actin filaments (396). Cap1/Cap2 localizes to actin patches, and strains carrying deletions in the CAP1 and/or CAP2 genes show depolarized patches and diminished cable staining (10, 11). The potential role of capping protein in regulating cable stability has not yet been addressed. However, recent studies show that capping protein plays an important role in driving actin polymerization to facilitate slow actin patch movements (Fig. (Fig.4)4) (174, 185). This effect may be explained by capping protein limiting the local concentration of free barbed ends of filaments to “funnel” available actin monomers to these barbed ends (i.e., those that are not capped) (287). This increases the rate of actin polymerization at these free ends, because the monomer pool is not depleted as rapidly and polymerization rates depend on monomer concentration.

Another important function that capping protein provides in actin networks is regulation of average filament length. By rapidly associating with free barbed ends of filaments, capping protein limits filament growth and thereby maintains a short average filament length. This is a critical property for the organization of dense actin networks comprised of many interconnected very short filaments, such as those found at the leading edge of motile cells, in the actin “comet tails” trailing intracellular vesicles and pathogens, or in yeast actin patches. Since capping protein is ubiquitously expressed and relatively abundant, the assembly of alternative actin arrays comprised of somewhat longer filaments (e.g., filopodia, stress fibers, and cables) requires that the filament length restrictions imposed by capping protein be actively bypassed. This is achieved by two mechanisms. First, capping protein can be regulated at the cell cortex by direct interactions with PIP2, which disrupts capping protein associations with barbed ends (332). Second, factors such as formins or Ena/VASP associate with the growing barbed ends of filaments (processive capping) and protect those growing ends from capping protein.

What is the mechanism for capping filament barbed ends? The recently solved crystal structure of vertebrate CapZ reveals that the two subunits, which have almost no similarity in primary sequence, fold into remarkably similar structures that intertwine (415). Each subunit has at its C terminus a short “tentacle” that protrudes from the main structural body of the protein, and mutational analysis shows that both tentacles contribute to capping in vitro. Further, mutations in the Cap1 tentacle cause cellular defects similar to those caused by the null mutations (186). Intriguingly, the Cap1 tentacle makes a much stronger contribution to capping than the Cap2 tentacle (186, 397). Further, the Cap1 tentacle is sufficient to cap filaments, albeit with far lower affinity than intact capping protein (186). Ultimately, a full biochemical understanding of the capping mechanism will require identification of the tentacle binding sites on actin, which have only been modeled to date and not determined experimentally (396).

Rapid Turnover of Actin Structures

Importance of actin turnover in vivo.

Although much research attention has been placed on defining the cellular mechanisms for assembling actin filaments within networks, the rapid disassembly of filaments is equally important. Only by maintaining actin networks in a state of dynamic flux, with individual filaments undergoing regulated turnover, are cells able to remodel their networks swiftly in response to internal and external cues (e.g., during cell motility and cytokinesis) and coordinate time-sensitive steps in complex actin-dependent processes (e.g., endocytosis). In addition, new growth of actin filaments relies on the rapid and steady turnover of older filaments, which is required to replenish the monomer pool. Thus, reduced rates of filament turnover deplete the monomer pool and limit new growth.

All three major actin structures in yeast (patches, cables, and rings) undergo rapid turnover, as demonstrated by the loss of all visible F-actin staining within 1 minute after treatment of cells with Lat-A (19, 369). As Lat-A is reported to block new filament growth but not actively disassemble filaments, this suggests that the filaments in patches, cables, and rings undergo rapid turnover. The importance of rapid actin turnover for cell function was first demonstrated in studies employing the cof1-22 conditional mutant (204). This mutation impairs the actin disassembly activity of Cof1 in vitro and dramatically reduces in vivo turnover rates of actin filaments in patches. This study showed a correlation between reduced rates of actin turnover and defects in endocytosis and cell growth. Similar results were reached in studies on act1-159, demonstrating a correlation between reduced rates of actin filament disassembly in vitro and reduced rates of actin turnover in vivo, accompanied by defects in endocytosis (27). Together, these studies strongly indicate that rapid turnover of actin is required for endocytosis and cell viability.

Most growth of actin networks in vivo occurs at the barbed ends of filaments, and disassembly is thought to occur primarily by dissociation of subunits from the pointed ends of filaments. However, the rate of dissociation of subunits from pointed ends is very low (0.3 to 0.8 subunits per s). Since rates of filament turnover in vivo are up to 100 times higher (428), this means that cells require active mechanisms for disassembling filaments. In yeast, as in other organisms, this is achieved through the coordinated activities of multiple actin-binding proteins working in concert. Below we describe the mechanisms and functions of each yeast actin-binding protein known to promote actin turnover. All of these proteins are highly conserved and, therefore, their mechanisms are likely to be conserved.


Budding yeast has a single essential cofilin gene (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 to accelerate actin filament depolymerization and turnover (204); however, the biochemical mechanism underlying this activity is not yet fully understood. Cofilin binds to actin filaments and monomers in vitro, in both cases preferentially interacting with ADP-bound versus ATP-bound actin (141). Cofilin induces filament fragmentation, which structural studies suggest is achieved by twisting of the filament to weaken lateral contacts between actin subunits (39, 241). As cofilin binds cooperatively to F-actin (140, 141), it may cluster at multiple sites along a filament, with nonuniform decoration leading to severing events. The other reported activity of cofilin, acceleration of subunit dissociation from filament pointed ends (51), could in principle also stem from filament twisting. Importantly, severing by cofilin has the potential to promote either assembly or disassembly of filaments depending on the availability of actin monomers in cells and how efficiently the new barbed ends generated by severing are capped.

What cellular function(s) requires the essential activities of yeast Cof1? Cof1 localizes to actin patches and has been shown to be required for endocytosis (204, 205, 246). Although cof1 mutants have not yet been studied using time-lapse imaging for defects in patch maturation, it seems likely that cof1-22 mutants will show defects in early slow patch movements, similar to capping protein mutants. Requirements at this phase of patch development seem to mirror those of other systems that depend on rapid actin polymerization and turnover, such as Listeria motility (221). In addition to its role in regulating actin turnover at patches, Cof1 may also play an unexpected role in driving actin cable dynamics. While Cof1 is detected specifically on actin patches in wild-type cells (246), it localizes to both patches and cables in aip1Δ cells (317). Further, recent functional studies show that cofilin (and Aip1) promotes the rapid turnover of actin cables and that aip1Δ mutations suppress the actin cable defects caused by deletion of TPM1 (276). Finally, a recent study showed that S. pombe Adf1 localizes to and functions in formation and maintenance of the actomyosin ring (264). Thus, Cof1 has the potential to regulate turnover of all the major actin structures in yeast cells: patches, cables, and rings.

One major open question about Cof1 is whether its activities are regulated in vivo. The ability of animal cofilins to interact with actin is regulated tightly by phosphorylation of a conserved serine residue in the N terminus, subject to control by a cascade of kinases and phosphatases (404). In yeast Cof1, an S4E (cof1-3) mutation of the analogous serine residue is lethal, suggesting that if phosphorylation occurred it would block actin binding and cofilin function, similar to what is seen in animal cells. However, no phosphorylation of endogenous Cof1 (isolated from exponentially growing cells) could be detected, which has led to the suggestion that Cof1 may not be regulated (205). One intriguing possibility, however, is that Cof1 may be down-regulated specifically when cells are not actively growing, such as in stationary phase. Another potential form of Cof1 regulation is pH change in the cytosol, as pH regulates the in vitro activities of cofilin/ADF proteins, including Cof1 (140, 141, 162), inhibiting filament disassembly activity at pH levels of <7.5. However, it remains uncertain whether pH is regulated locally in yeast cells and, therefore, whether this regulatory mechanism contributes to Cof1 function in vivo. Finally, the activity of ADF/cofilins is down-regulated by direct interactions with PIP2 (250, 272), which may have important implications for regulation of Cof1 at cortical sites, such as actin patches.


Aip1 is an important binding partner and cofactor of cofilin 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-interacting factor through biochemical affinity (275), two-hybrid (317), and genetic (163) approaches. Together, Aip1 and cofilin promote net actin filament disassembly in vitro, and aip1Δ and cof1 mutations show genetic interactions (7, 244, 275, 317). Like cofilin, Aip1 localizes to actin patches (317), where it has a demonstrated role in promoting rapid actin turnover (276).

Elucidation of the Aip1 mechanism has seen much recent progress. Aip1 caps the barbed ends of cofilin-severed filaments 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 β-propellers linked by a short hinge domain (Fig. 13A) (245, 384). Mutagenesis studies have defined two actin-binding surfaces on Aip1 that are critical for its functions in vitro and in vivo, one located on each β-propeller (66, 243, 245, 276). The cofilin-binding surface lies in the hinge region of Aip1 located between 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 promotes filament disassembly specifically in the presence of cofilin, this C-terminal actin-binding site may recognize a cofilin-induced conformational state of F-actin. These properties tailor Aip1 for capping cofilin-severed filaments and, indeed, mutation of the C-terminal actin-binding site abolishes Aip1's cofilin-dependent activity. Taking these observations together, a model emerges for the Aip1 mechanism and function. Cofilin decorates the sides of filaments and helps recruit Aip1, which binds directly to F-actin and cofilin. Together, cofilin and Aip1 induce filament severing to produce short fragments of F-actin (less than 50 nm) capped at their barbed ends by Aip1. These fragments disassemble rapidly by the dissociation of subunits from their pointed ends.

FIG. 13.
Regulation of actin filament turnover by a network of interacting proteins. (A) Schematics of proteins discussed in the actin turnover section of the text. Abbreviations: β-propeller, forms β-propeller tertiary structure composed of WD ...

Until recently, Aip1 and cofilin were thought to promote actin turnover specifically at patches, owing to their localization to patches. However, as discussed in above (see “Actin cable dynamics and turnover”), cofilin and Aip1 have recently been shown to promote rapid turnover of cables (276). The rapid nature of the cofilin-Aip1 mechanism of disassembling filaments may explain why in wild-type cells cofilin and Aip1 are not detected on cables. Cables are faint to begin with, and if only a small percentage of filaments are decorated with cofilin, it would be difficult to detect. In contrast, in aip1Δ cells, severing by cofilin produces uncapped barbed ends that undergo new polymerization, consistent with cable thickening and detection of cofilin on cables. This also explains why thickened cables decorated with cofilin are observed in the cof1-19 strain, a mutant that disrupts barbed end capping by Aip1 (317).


Srv2 was initially identified in yeast as a suppressor of the activated rasV19 allele (Srv2) and as an adenylyl cyclase-associated protein (CAP) (103, 107). The yeast protein was named Srv2, but homologues in other species are called CAP. Yeast Srv2 interacts with the adenylyl cyclase complex through a binding site in its N terminus, whereas the C terminus of the protein binds to actin monomers and plays a central role in regulation of actin dynamics in cells (Fig. 13A) (112, 117, 180, 237, 424). The adenylyl cyclase-binding activity does not appear to be conserved in animals and plants, but its role in regulating the actin cytoskeleton is conserved in all Srv2/CAPs tested so far (158). Deletion of the SRV2 gene in yeast causes defects in cell growth and actin organization, and expression of CAP homologues from diverse species rescues these defects. In addition, srv2Δ defects are suppressed partially by overexpression of profilin, suggesting a relationship between their in vivo functions (385). Until recently, the specific function performed by Srv2/CAP in the actin cytoskeleton was unknown.

Recent studies have revealed that Srv2/CAP promotes the rapid turnover of actin networks by recycling newly depolymerized actin monomers and cofilin (23, 31, 237, 249). After cofilin promotes filament disassembly, it remains bound to ADP-actin monomers and inhibits nucleotide exchange (ATP for ADP) (36, 205, 269). This leads to the accumulation of a pool of cofilin-bound ADP-actin monomers, blocked from returning to an ATP-bound assembly-competent state. Therefore, cells require a mechanism to displace cofilin from ADP-actin monomers. Classical models depict profilin performing this function through competition (295). However, profilin fails to compete effectively with cofilin for binding to ADP-G-actin, suggesting that a “middle man” is required to facilitate exchange of actin monomers from cofilin to profilin. Srv2/CAP has been demonstrated to perform this function in vitro (23). The C terminus of Srv2/CAP binds to and promotes nucleotide exchange on actin monomers. Further, full-length Srv2 catalyzes the conversion of cofilin-bound ADP-actin monomers to profilin-bound ATP-actin monomers. Thus, Srv2/CAP promotes actin turnover by (i) recycling cofilin for new rounds of filament disassembly 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 mechanism underlying these effects (237). The C terminus of Srv2 has an ∼100-fold-higher affinity for ADP-G-actin (Kd = 0.02 μM) than ATP-G-actin (Kd = 1.9 μM), which enables it to compete for ADP-G-actin binding with cofilin (Kd = 0.02 to 0.1 μM cofilin) (36, 51, 237, 379). Further, the N terminus of Srv2 may assist in these events, as it binds specifically to cofilin-G-actin complexes 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 terminus of Srv2 has the ability to weaken cofilin-ADP-actin monomer interactions. Following transfer of ADP-G-actin from cofilin to Srv2, rapid nucleotide exchange occurs on actin. However, Srv2-bound ATP-actin monomers are not competent for addition onto filament barbed ends (237). This means that another handoff is required (to profilin). Addition of profilin to Srv2-bound G-actin releases monomers for addition onto barbed ends, suggesting a transfer has occurred (237). Thus, Srv2/CAP catalyzes conversion of actin monomers from the ADP- to ATP-bound state and, like many enzymes, Srv2 has a significantly 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, is that 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 ∼15 by 30 nm long as determined by rotary shadowing EM of purified complexes (23). The complex is comprised entirely of two proteins, Srv2 and actin, present at a 1:1 molar stoichiometry, suggesting a heterododecamer (assembled from six Srv2 and six actin molecules). Each Srv2 molecule in the complex can interact with G-actin and at least three actin-binding proteins: cofilin, profilin, and Abp1. Binding of cofilin and profilin may promote the handoffs described above. Binding of Abp1 is required for Srv2 localization to yeast actin patches (214) and tethers Srv2 complex to actin filaments in vitro (23). Since the Abp1-binding site on Srv2 is located in its C terminus, proximal to the G-actin-binding domain, it will be interesting to learn whether Abp1 also affects Srv2 activity. Taken together, these interactions suggest that the Srv2 complex may serve as a molecular hub to facilitate dynamic interactions among multiple proteins in promoting actin turnover. This concept raises important questions about the structure and physical properties of the complex and how they influence function. For instance, what is the nucleotide-bound state of the actin subunits bound to the complex? Do these subunits freely exchange with actin monomers in solution? How are Srv2 and actin molecules arranged within the complex? Is multimerization of Srv2 required for its activities and in vivo functions?


Profilin is a small (∼15-kDa), ubiquitous, abundant actin monomer-binding protein, the biochemical activities of which have long been studied (409). In all organisms examined, mutation of profilin is lethal and/or causes severe defects in actin organization and cell morphology. In S. cerevisiae, a deletion mutant (pfy1Δ) and point mutations that impair actin binding (e.g., pfy1-4) show that profilin is critical for cell growth, morphology, and actin organization (134, 410).

Association of profilin with actin monomers (Fig. (Fig.8)8) has four known effects of potential importance in vivo. First, it suppresses spontaneous actin assembly by sterically hindering interactions between actin monomers required for the formation of polymerization intermediates (295). Second, it restricts actin monomer addition to filament barbed ends, blocking addition to pointed ends (297). Third, it can increase the rate of elongation at filament barbed ends capped by formins (195, 321). Fourth, it accelerates nucleotide exchange (ATP for ADP) on G-actin (242). In biomimetic assays for cell motility that employ purified proteins, profilin becomes rate limiting for actin assembly, consistent with its proposed function in accelerating actin turnover (206, 221, 230). However, it remains unclear which activities of profilin are required in vivo.

Synthetic lethal interactions between pfy1-4 and cof1-22, srv2Δ, and act1-159 are consistent with profilin promoting actin turnover in vivo. However, it is important to note that direct evidence for this proposed function is still lacking, as actin turnover rates have not been measured for profilin mutants in vivo. One hypothesis is that profilin's ability to promote nucleotide exchange on G-actin is critical for its proposed turnover function. This is supported by the observation that cellular defects caused by the pfy1-4 allele (impaired in binding actin) are suppressed by act1-157, an allele with an increased intrinsic rate of nucleotide exchange (410). This study concluded that profilin has a critical role in vivo in accelerating actin monomer recycling. However, other observations suggest that profilin may contribute to actin turnover by a mechanism other than promoting nucleotide exchange on G-actin. First, S. cerevisiae profilin increases nucleotide exchange on G-actin by only ∼3-fold in vitro, orders of magnitude less than human and birch pollen profilin (88). Second, there are no mutants yet available that impair nucleotide exchange without disrupting actin binding. Thus, it is not possible to separate effects of profilin on nucleotide exchange from its other effects, such as promotion of 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 pfy1Δ yeast cells (62); this raises the possibility that acceleration of nucleotide exchange may not be a critical function of profilin in yeast.

What turnover function might profilin perform beyond accelerating nucleotide exchange on G-actin? Recent studies (see “Srv2/CAP” above) suggest that profilin plays a key role along with Srv2/CAP in facilitating the rapid conversion of actin monomers (from a cofilin-bound ADP-G-actin state to an assembly-competent profilin-bound ATP-G-actin state). In this proposed mechanism, profilin binds to G-actin following nucleotide exchange. This is consistent with (i) profilin having very low affinity for ADP-G-actin and significantly higher affinity for ATP-G-actin (383) and (ii) profilin failing to compete with cofilin for ADP-G-actin binding (23). In cells, profilin may first bind to ATP-actin monomers while they are still associated with Srv2/CAP, just after they have undergone nucleotide exchange (see above). Srv2/CAP has a weak affinity for ATP-G-actin, which facilitates the handoff to profilin (237). In addition, since Srv2-bound actin monomers cannot be added to filament barbed ends, an additional handoff is required to factors such as profilin that readily promote barbed end assembly. This model is supported by the ability of profilin to siphon ATP-actin monomers from Srv2/CAP and make them available for barbed end growth (237). It is also supported by physical interactions between profilin and Srv2/CAP that may facilitate monomer handoffs (32, 82). Thus, the loss of profilin would stall this mechanism and rapidly deplete the assembly-competent pool of actin monomers.

Another point to consider is that acceleration of nucleotide exchange by profilin requires its interaction with actin but not proline-rich sequences, yet mutations in profilin that impair polyproline binding (without affecting actin binding) cause severe defects in cell growth and actin organization (134, 225, 410). This suggests that profilin function in vivo depends on interactions with both actin and polyproline targets, indicating that profilin must have critical functions beyond nucleotide exchange on actin. What is the other function of profilin in vivo? Profilin targets actin monomers to the growing barbed ends of filaments associated with Ena/VASp and formins. In S. cerevisiae, which does not express Ena/VASP homologues, profilin is believed to target actin monomers to barbed ends of filaments that the formins Bni1 and Bnr1 assemble into cables and the actomyosin ring. Profilin binding to the proline-rich formin 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 in vivo, profilin mutants impaired in binding polyproline display severe defects in actin cable assembly and polarized cell growth (134, 225, 410).


Twinfilin is a widely conserved 40-kDa protein comprised of two tandem ADFH domains (Fig. 13A). The founding member of the twinfilin family (Twf1) was identified in S. cerevisiae by mining the genome database for cofilin homologues and by isolation from crude extracts on F-actin affinity columns with other actin-binding proteins, including capping protein (125). Initial biochemical characterization of Twf1 showed that it is an actin monomer-sequestering protein. Although twf1Δ mutants showed normal cell growth and morphology, their actin patches were slightly brighter, consistent with actin turnover defects, and recent work has demonstrated that loss of TWF1 reduces rates of actin patch turnover (252). twf1 mutants also are synthetic lethal with cof1 and pfy1 mutants (125), consistent with a role in promoting actin turnover.

Biochemical analyses have left the precise role of Twf1 in actin turnover somewhat enigmatic. Twinfilin sequesters actin monomers (125), with a strong binding preference for ADP-G-actin (Kd ∼ 0.05 μM) over ATP-G-actin (Kd ∼ 0.5 μM) (273). While each ADFH domain of twinfilin is capable of binding G-actin, the C-terminal ADFH domain binds to G-actin with affinity similar to that of intact twinfilin, suggesting that the N-terminal ADFH domain may have an alternative function. Twf1 also binds directly to capping protein (Cap1/Cap2) and requires capping protein for its localization to actin patches in vivo and for genetic functions shared with cofilin (101, 285). However, the Twf1-capping protein interaction does not affect the capping activity of Cap1/Cap2 or the monomer-sequestering activity of Twf1. Thus, two discrepancies regarding Twf1 function have been the following: how do Twf1-actin interactions promote actin turnover in vivo, and what function is served by the Twf1-capping protein interaction?

Two recent studies have shed new light on these issues. In one study, mammalian twinfilin was demonstrated to cap filament barbed ends; however, this activity was not conserved in yeast Twf1 (143). In a second study, yeast Twf1 was found to bind to and sever actin filaments at low pH (252), an intriguing result given the known pH dependence of ADF/cofilin activities on F-actin (24, 85, 162). Further, capping protein directly inhibits the filament-severing activity of Twf1 (252), representing the first biochemical function ascribed to this physiological interaction. Presently, it is difficult to explain the in vivo significance of the low-pH dependence of filament-severing activity in vitro. One possibility is that posttranslational modifications of Twf1 in vivo enable it to sever filaments at more neutral pH levels. Alternatively, Twf1-severing activity may be regulated by pH fluctuations in vivo.

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


Over the past decade, intense research efforts have shown that the three most prominent actin structures (patches, cables, and rings) are all highly dynamic, with patches and cables assembling and turning over in less than 60 seconds. Further, numerous genetic, two-hybrid, and biochemical studies have brought us close to having a complete list of actin-associated proteins involved in these processes. For many, their activities on actin have been determined (Table (Table1)1) and their atomic structures now are available (e.g., Abp1, the Arp2/3 complex, cofilin, coronin, profilin, Sac6, Srv2/CAP, and twinfilin). All of this sets the stage for a detailed mechanistic dissection of the elegantly choreographed events in actin network assembly and function.

One key future challenge is to develop new biochemical strategies for defining the activities of actin-associated proteins in cellular mixtures. To date, the effects of actin-binding proteins on actin have been studied primarily using in vitro assays containing one or two purified actin-binding proteins and actin, and often these components are not present at concentrations or molar ratios found in vivo. However, the activities of actin-binding proteins can be affected strongly by their in vivo binding partners (e.g., Crn1-Arp2/3, Aip1-cofilin, and Srv2/CAP-cofilin). Most actin-binding proteins do not function alone but rather in stable and/or dynamic complexes. Further, actin-binding proteins must compete for limited binding sites on actin, which can greatly influence their ability to affect actin dynamics and organization. Thus, the activities of actin-binding proteins should be examined in complex mixtures that more closely reflect physiological conditions. Some of these complexes can be isolated from yeast cells in native form; others can be “built up” from purified proteins with known interactions. This also emphasizes the need to define local cellular concentrations and in vivo molar ratios of actin-binding proteins, as has been done recently for some actin-binding proteins in fission yeast (413).

A second future challenge is to understand how cellular signals regulate the activities of actin-binding proteins to alter actin filament assembly, organization, and turnover. Live cell imaging studies have revealed that the assembly and disassembly of yeast actin networks (patches, cables, and rings) are subject to exquisite spatial and temporal regulation. Further, many signaling molecules have been implicated in regulating actin cytoskeleton organization and function (e.g., Pkc1, Ark1/Prk1, Ste20/Cla4, Cdc28, and mitogen-activated protein kinases). However, in the vast majority of cases the actin-binding protein targets of these signaling pathways are unknown, and therefore the mechanisms of regulation are unclear. One of the few examples of direct regulation recently demonstrated is phospho-regulation of Pan1 NPF activity by Prk1 kinase (371). Additionally, Rho GTPases directly regulate actin-binding proteins such as Bni1 and Bnr1 and control Ste20/Cla4 kinases and the polarisome scaffold protein Bem1 (170). More studies of this nature are needed, as these examples likely represent only the tip of the iceberg for signaling molecules governing the activities of actin-binding proteins.


We are very grateful to all who contributed to the assembly and editing of this review. We are especially grateful to Jacqueline Hendries for her devoted assistance during the early stages and to the following people for their generous help in editing the review prior to submission: S. Buttery, F. Chang, K. Daugherty, M. Gandhi, K. Kono, S. Martin, K. Okada, D. Pruyne, A. Rodal, and I. Sagot. We also thank the reviewers for valuable suggestions and D. Amberg, D. Botstein, A. Bretscher, M. Eck, J. Pringle, L. Pon, A. Rodal, and M. Rosen for their permission to reproduce original images. Finally, we apologize for the omission of important contributions made to this field that we may have overlooked in writing this review.

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


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