Virtually all eukaryotic cell types have morphologies that are uniquely tailored to their physiological functions. The immense variation in cell shape depends crucially on an underlying network of dynamic, interconnected actin and microtubule polymers. The dynamic assembly and turnover of these filamentous networks is used to direct cell polarity and to facilitate membrane and organelle traffic, cell adhesion, chromosome segregation, cell migration and cell division.

To construct these intricate fibrous arrays, cells make use of a palette of proteins that bind to cyto skeletal polymers and work in concert to organize them into higher-order force-generating structures. The task of assembling actin and microtubule polymers de novo requires active mechanisms, as there are abundant factors in cells that inhibit spontaneous polymer formation. These inhibitory factors include proteins that buffer free actin sub units (such as profilin and thymosins) and tubulin subunits (such as stathmin), and factors that cap polymer ends and sever or depolymerize polymers. To build new actin and microtubule polymers, cells deploy specialized proteins that catalyse polymer nucleation and elongation, protect growing polymer ends and attach to the sides and/or ends of polymers to stabilize them against disassembly.

Recent work has shown that the formin family of pro-teins (which is conserved in plants, animals and fungi) has a central role in catalysing actin polymer assembly and in stabilizing microtubules1,2. Various other proteins that are capable of stimulating actin assembly are found in cells, including the Arp2/3 complex, spire, cordon-bleu (COBL), leiomodin (LMOD), and junction-mediating and -regulatory protein (JMY).3,4. However, formins

might be unique in their ability to directly regulate both actin filaments and microtubules. Furthermore, only formins show a clearly established and robust ability to both nucleate actin polymers and dramatically accelerate polymer elongation.

Formins are large (120–220 kDa), multidomain pro-teins that interact with many binding partners to perform their functions (TABLE 1). Fungal species typically have 2 or 3 formin genes, whereas mammals have 15 and some plant species have more than 20 (REfs 5,6,7). The potent activities of formins on actin and micro tubule dynam-ics have been harnessed to the assembly of the diverse cytoskeletal structures that are required in a range of cell-ular functions, including cell morphogenesis, adhesion, division and motility (fIG. 1). Formins are also implicated in a growing number of diseases (BOX 1).

In this review, we first summarize formin structure and activities and then we describe regulatory control points and mechanisms used to govern formin activities.

Deconstructing forminsAlthough no three-dimensional (3D) structure has yet been reported for any intact formin, a working model for their general architecture can be assembled from the crystal structures of formin fragments (fIG. 2). In this model, formins are depicted as dimers, as recent biophysical data has confirmed that the purified full-length formins mouse diaphanous 1 (mDia1; also known as DIAPH1) and mDia2 (also known as DIAPH3), and yeast Bni1 and Bni1-related protein 1 (Bnr1) are dimeric (B.G., unpublished observations). Below, we describe the general domain layout of animal, fungal and plant formins.

Rosenstiel Basic Medical Science Research Center, Brandeis University, Waltham, Massachusetts, 02454, USA.Correspondence to B.G. e‑mail: goode@brandeis.edudoi:10.1038/nrm2816 Published online 9 December 2009

Unleashing formins to remodel the actin and microtubule cytoskeletonsMelissa A. Chesarone, Amy Grace DuPage and Bruce L. Goode

Abstract | Formins are highly conserved proteins that have essential roles in remodelling the actin and microtubule cytoskeletons to influence eukaryotic cell shape and behaviour. Recent work has identified numerous cellular factors that locally recruit, activate or inactivate formins to bridle and unleash their potent effects on actin nucleation and elongation. The effects of formins on microtubules have also begun to be described, which places formins in a prime position to coordinate actin and microtubule dynamics. The emerging complexity in the mechanisms governing formins mirrors the wide range of essential functions that they perform in cell motility, cell division and cell and tissue morphogenesis.

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Polyproline tractA short protein motif, found in many actin regulatory scaffold proteins, that typically contains five or more tandem proline residues and binds profilin or sH3 domains.

Barbed endThe rapidly growing end of an actin filament, so-called because of the arrowhead pattern created when myosin binds. The slowly growing end is called the pointed end.

Formin polypeptides can be divided into two major functional regions (fIG. 2a): the amino-terminal ‘regu-latory’ region, which typically governs in vivo local-ization and can influence the activities of the carboxy terminus, and the ‘active’ region, which stimulates actin assembly and, in some formins, interacts with micro-tubules. The C terminus of many formins includes a Dia autoregulatory domain (DAD), which can, in some cases, mediate auto inhibition through interactions with the n terminus.

The C-terminal active region consists of the formin homology 1 (FH1) and FH2 domains. The FH1 domain is predicted to be rope-like, based on a lack of secondary

structure, and it contains binding sites for profilin–actin complexes. Profilin is a ubiquitous actin monomer- binding protein with separate binding sites for mono-meric actin (also called globular actin or G-actin) and poly proline tracts8–10. Profilin is associated with most actin monomers in cells11 and, therefore, profilin–actin com-plexes are the predominant substrate for actin assem-bly in vivo. Interactions between profilin and the FH1 domain are crucial for the recruitment of actin mono-mers to the active region12–15. The adjacent FH2 domain forms a head-to-tail doughnut-shaped dimer that en circles the barbed end of the actin filament. In mDia1 or mDia2, the FH2 domain, plus flanking sequences,

Table 1 | Binding proteins regulating formin localization and activity

Protein* Formin target* Organism and/or cell type Function Refs

Profilin All tested All cell types Recruits actin monomers to the FH1 domain to accelerate elongation 53

Bud6 Bni1 Saccharomyces cerevisiae Nucleation cofactor for Bni1, binds to the DAD and promotes the assembly of actin cables

41,149

Fus3 Bni1 S. cerevisiae Phosphorylates and localizes Bni1 to the tips of mating cells 95

Rho1 Bni1 S. cerevisiae Required for Bni1 localization to the bud neck and bud cortex 97,150

Spa2 Bni1 S. cerevisiae Helps localize Bni1 to the bud cortex 103,151

Bud14 Bnr1 S. cerevisiae Displaces the Bnr1 FH2 domain from growing barbed ends of filaments and regulates actin cable architecture

89

Spire CAPU Drosophila melanogaster oocytes

Synergizes with CAPU to assemble actin meshworks in vivo and is thought to inhibit CAPU in vitro

44,45

Cdc15 Cdc12 Schizosaccharomyces pombe Binds directly to the amino terminus of Cdc12 and is required for the assembly of the cytokinetic actin ring and for cell division

152

Bud6 For3 S. pombe Binds to the DAD and helps localize For3 to cell tips and is required for actin cable assembly

42,153

Cdc42 For3 S. pombe Helps localize For3 to cell tips, is required for actin cable assembly and interacts genetically with Bud6

42

Tea4 For3 S. pombe Helps localize For3 to cell tips, is required for actin cable assembly and is the closest homologue of S. cerevisiae Bud14 by sequence

104

ABI1 mDia1 Epithelial, melanoma and HeLa cells

Helps localize mDia1 to lamellipodia, filopodia and cell adhesions 105,147

CLIP170 (CLIP1)

mDia1 (DIAPH1) Macrophages Binds the FH2 domain and helps localize mDia1 to sites of phagocytosis

17

Gα12/13

mDia1 Fibroblasts Helps localize mDia1 to the leading edge of migrating cells 154

IQGAP1 mDia1 Fibroblasts and macrophages Binds to the DID and is required for mDia1 localization to the leading edge and to phagocytic cups

93

RHOA mDia1 Epithelial cells Required for mDia1 localization to adherens junctions and partially activates mDia1 from autoinhibition

26,100

RHOB mDia1 Melanoma cells Helps localize mDia1 to endosomes 155

RHOB mDia2 (DIAPH3) Fibroblasts Required for mDia2 localization to endosomes 25

RIF (RHOF) mDia2 Fibroblasts Helps localize mDia2 to filopodial tips 156

DIP (WISH, NCKIPSD)

mDia2 HEK and HeLa cells Inhibits mDia2 FH2, suppresses filopodial protrusion and induces membrane blebbing

88

ROCK1 FHOD1 HeLa cells Binds the FH2 domain and phosphorylates and activates FHOD1 to promote membrane remodelling

39,40

α-catenin FMN1 Epithelial cells Helps localize FMN1 to adherens junctions and is required for FMN1-dependent actin assembly at cell adhesion sites

106

Cdc42 FMNL1 (FRL1) Macrophages Helps localize FMNL1 to the cell cortex 84

ABI1, Abelson interactor 1; Bnr1, Bni1-related protein 1; Bud, bud site selection protein; CAPU, Cappuccino; CLIP170, cytoplasmic linker protein 170; DAD, Dia autoregulatory domain; DID, Dia inhibitory domain; DIP, Dia-interacting protein; FH, formin homology; FHOD1, FH1/FH2 domain-containing protein 1; FMN1, formin 1; FMNL1, FMN-like protein 1; For3, formin 3; IQGAP1, IQ motif-containing GTPase activating protein 1; mDia, mouse diaphanous; RIF, Rho in filopodia; ROCK, Rho-activated kinase; Tea4, tip elongation aberrant protein 4. *Alternative protein names are provided in brackets.

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Nature Reviews | Molecular Cell Biology

Nucleus

Lamellipodium

Filopodiuma b

dc

Stress fibre

Microtubule

Phagocytic cup

Endosomaldynamics

Adherens junction

0 5 10 15 20Size (µm)

2.5 Sec5 Sec

7.5 Sec10 Sec

Transport Transport

Actinfilament

Actincable

Cytokinetic actinring

Actin filamentFormin

Microtubule plus end tracking proteinOne of a group of proteins that are enriched at the fast growing (plus) ends of microtubules and that influence microtubule dynamics and/or link microtubule plus ends to other cellular structures.

Coiled-coil domainA structural domain that can mediate protein oligomerization. Coiled coils are helices that are assembled from repeat sequences of seven amino acids (heptads) and twist around each other to form a supercoil.

Armadillo repeatA folded helical structure encoded by a conserved sequence that was first identified in the D. melanogaster protein Armadillo. This domain is conserved in animals and higher plants and is found in various signalling and cytoskeletal proteins.

also binds directly to microtubules and interacts with three microtubule plus end tracking proteins: end-binding p rotein 1 (eB1; also known as MAPRe1), adenomatous polyposis coli (APC) and cytoplasmic linker protein 170 (CLIP170; also known as CLIP1)16–18. FH2 is the most conserved domain in formins. Other formin sequences and domains vary considerably, particularly between plant and animal or plant and fungal formins.

Phylogenetic analyses have classified the 15 distinct mammalian formin genes into 7 subfamilies5,19 (TABLE 2). In most subfamilies, and in fungal formins, the n terminu s consists of a Rho-binding domain (RBD), a Dia inhibitory domain (DID) and a dimerization domain, which is some-times followed by a coiled-coil domain (fIG. 2a). The DID comprises tandem armadillo repeats and, in some formins, binds to the C-terminal DAD20,21,22. The DAD consists of a short α-helix23 and flanking basic residues24,25. For some formins, DAD–DID interactions strongly inhibit actin assembly by the FH1 and FH2 domains, although the structural basis of this inhibition is not understood. In specific cases, autoinhibition can be relieved by Rho binding to the RBD26,27, but increasing evidence points to the involvement of additional cell ular factors in formin activation (see below). In addition, some formins appear to activate their Rho binding partners, raising the possibility of reciprocal activation28,29.

whereas the n termini of most animal and fungal formins follow the general domain layout described above, there is a high degree of variability in the RBD and possible ligands. Furthermore, some formins lack a recognizable RBD, and the n terminus of one formin subfamily (neuronal-specific delphilin) lacks RBDs,

DIDs and dimerization domains and, instead, contains one or two PDZ domains30,31. One PDZ domain interacts with the δ2 glutamate receptor subunit 31, which co localizes with delphilin at postsynaptic nerve terminals and is required for normal synaptic function30.

All known plant formins lack RBDs, DIDs and dimer-ization domains and, instead, contain either transmem-brane or PTEN domains. Conventional PTen domains have lipid phosphatase activity, but the sequence of plant formin PTen domains suggests that they might be cata-lytically inactive and have non-enzymatic functions such as localization to membranes6. These differences in plant formins suggest that they might directly link membranes to actin assembly, which could underlie their requirement in cell wall and cell shape remodelling32.

Formin activities and mechanismsFormins govern two distinct phases of actin assem-bly: nucleation and elongation. These properties were revealed by two complementary in vitro assays for meas-uring actin dynamics — pyrene–actin assays and total internal reflection fluorescence microscopy (TIRF) (BOX 2). The biochemical effects of formins on actin are strong, typically requiring only 5–200 nM formin for robust nucleation and/or elongation activity in vitro. within this range however, formin potency can vary greatly. It is yet to be determined whether such differences reflect distinct functional requirements in vivo. Interestingly, all formin biochemical activities to date have been defined using recombinant or overexpressed proteins. Thus, the eventual purification of endogenous formins offers the potential to expose new activities and regulation.

Actin nucleation. The FH2 domain is sufficient to catalyse nucleation in vitro from purified actin mono-mers14,33. Although the FH2 domain lacks a detectable binding affinity for monomers, it binds with high affinity (with a low nanomolar Kd) to the barbed ends of actin filaments. These interactions depend on the ability of the FH2 domain to dimerize34,35. each functional half of the FH2 dimer is called a hemi-dimer or ‘bridge’ and contains two f-actin-binding sites35,36 (fIG. 2).

Precisely how formins catalyse actin nucleation has remained elusive. The rate-limiting step in de novo actin polymerization is the self-association of monomers into short-lived polymerization intermediates (dimers and trimers). when these kinetic barriers are overcome, a tetramer is formed. The tetramer is a stable seed for polymerization, which continues for as long as the actin monomers are above the critical concentration37. All other known nucleators and/or their cofactors bind to and recruit multi ple actin monomers to stimulate polymer for-mation3. The FH2 domain, however, does not bind actin monomers. Therefore, it has been hypo thesized that the FH2 domain binds and stabilizes spontaneously formed actin dimers and trimers to stimulate poly merization, and this is supported by kinetic modelling33. Although this explains how the FH2 domain nucleates actin assembly in vitro, it does not explain how formins nucleate actin assembly in vivo, where the monomer pool is associated with profilin and thymosins, which inhibits spontaneous

Figure 1 | cellular structures and functions controlled by formins. a | Formins play multiple roles in migrating cells. They catalyse the assembly of actin filaments (red) to help drive the formation of filopodial processes146, lamellipodial sheets147 and stress fibres81. Formins also stabilize microtubules (blue) to help steer cell migration16,65. b | Some formins (green) move rapidly with the growing ends of actin filaments in vivo at rates of approximately 2 microns per second49. c | Formins are also required for the assembly of actin-rich adherens junctions106, which contain cell adhesion and adaptor proteins, and for normal rates of phagocytosis84 and endosomal movement115. d | In budding yeast, formins are essential for the assembly of actin cables that direct membrane transport, and for the assembly of cytokinetic actin rings79.

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PDZ domain(Postsynaptic-density protein of 95 kDa, Discs large and Zona occludens-1 domain). A protein-interaction domain that often occurs in scaffolding proteins and is named after the founding members of the family.

PTEN domain(Phosphatase and tensin domain). A conserved lipid protein phosphatase domain that has been extensively characterized in the tumour suppressor protein PTEN. Many PTEN domain-containing proteins regulate cytoskeletal organization.

Total internal reflection fluorescence microscopyA microscopy technique that enables the real-time visualization of fluorescently labelled molecules in a thin region of a specimen, greatly reducing background, and that has enabled actin dynamics to be studied at the single filament level.

self-association. However, specific conditions in living cells, including rapid filament dis assembly, might trans-iently elevate the local concentration of free G-actin to promote formin-mediated nucleation events38.

In many other cases, nucleation by formins in vivo might require actin monomer-binding sequences out-side of the FH2 domain and/or nucleation cofactors that bind actin monomers. Although FH1 recruits actin monomers through profilin binding, it does not promote nucleation15. However, sequences on the C-terminal side of FH2 might directly and/or indi-rectly recruit actin monomers to promote nucleation. Indeed, the Saccharomyces cerevisiae polarity factor bud site selection protein 6 (Bud6) binds to actin mono-mers and to the formin Bni1 (REf. 34) to dramatically increase Bni1 nucleation efficiency (A.G.D and B.L.G, un published observations)34. Bud6 shares sequence homology with a non-kinase domain of Rho-activated kinase (ROCK1), a mammalian ser/Thr kinase impli-cated in Rho-mediated actin organization that directly regulates the formin FH1/FH2 domain-containing protein 1 (FHOD1) in vivo39,40. Therefore, the use of nucleation cofactors by formins might be conserved. Interestingly, Bud6 binds to the autoregulatory DAD of Bni1 (REfs 41,42), suggesting that the DAD has multiple roles, regulating both autoinhibition and recruitment of actin monomers to promote nucleation. similarly, sPIRe1, an actin monomer binding protein that nucle-ates actin assembly in vitro, binds to a motif at the

C-terminal side of the FH2 domain in formin 2 (FMn2) that is conserved in other FMn proteins43. spire binding to FMn has been suggested to either inhibit nucleation or to have no effect44,45. Together, these findings suggest that sequences at the C-terminal side of FH2 can attract ligands with the ability to either positively or negatively influence nucleation activity.

Actin elongation. Once a filament is nucleated, the dimeric FH2 domain moves processively with the grow-ing barbed end, shielding it from capping proteins, while permitting the rapid addition of new subunits46–48. The rate of FH2 movement matches the rate of actin sub-unit addition, which can exceed 100 subunits per sec-ond48–50. This rapid movement might be achieved by FH2 hemi-dimers alternating their contacts with two actin subunits exposed at the barbed end. each trans-ition or ‘step’ that the FH2 domain makes might be trig-gered by the addition of the previous actin subunit1. The flexible linker connecting the FH2 hemi-dimers is thought to play a crucial part in these events by accom-modating drastic spatial rearrangements of the hemi-dimers with respect to each other, which is predicted to occur with each step. To date, only the actin remodel-ling protein ENA/VAsP has been shown to have a similar ability to move with and protect a growing barbed end. unlike formins, this ability requires the clustering of enA/VAsP oligomers to a surface, suggesting key mechanistic differences, but some aspects of their mech-anisms might be related3,51. An important distinction between the processive movement of formins and classic motor proteins is that formins do not bind or hydrolyse nucleotides. One study suggested that the energy for FH2 movement is derived from profilin-stimulated ATP hydrolysis on actin52. However, sub-sequent studies showed that FH2 movement does not require profilin or ATP hydrolysis and, instead, might be coupled to the release of free energy accompanying actin polymerization15,53,54.

The rate of elongation at FH2-capped barbed ends is influenced greatly by interactions of the adjacent FH1 domains with profilin–actin. Through these inter-actions, formins can accelerate actin subunit addition at FH2-capped ends by up to 19-fold compared to rates of diffusion-limited growth at free barbed ends50,52,53. The mechanistic basis for the formin ‘gas pedal’ effect is not yet understood, but might result from FH1 domains increasing the local concentration of actin monomers near the barbed end and/or pre-orienting actin mono-mers for efficient addition to the barbed end. speed of elongation varies greatly among formins50,53 and was shown recently to depend on the pairing of specific profilin isoforms with FH1 domains and specific FH1 domains with FH2 domains55. However, the precise structural parameters in the FH1 and FH2 domains that specify elongation rate are yet to be determined. In addi-tion, it is unknown whether formins with distinct speeds are tailored to assemble specific actin structures in vivo, and/or whether combinations of formins with different speeds are co-employed to construct actin networks comprising filaments of different lengths.

Box 1 | Formins in disease

Formins are linked to the progression of cancer and other diseases, which probably stems from their instrumental roles in cell division and migration. In human cells, overexpression of formin-like protein 1 (FMNL1; also known as FRL1) is linked to leukaemia124 and non-Hodgkin lymphoma125, and elevated expression of FMNL2 (also known as FRL2) is associated with metastasis of colorectal cancer cells126. Members of the diaphanous (Dia) subfamily of formins promote the formation of actin-rich invadipodia in aggressive breast cancer cells127, and aberrant activation of mouse Dia1 (mDia1; also known as DIAPH1) increases the protrusive activity of carcinoma cells80. mDia1 also directly activates leukaemia-associated Rho-GEF (LARG; also known as ARHGEF12) to increase RHOA activity29, which is associated with various cancers128. Furthermore, downregulation of mDia1 triggers Rho-associated protein kinase (ROCK)-dependent invasiveness in melanoma cells and is linked to haematopoietic disorders129,130. Genome-wide studies have identified formin gene mutations in brain and pancreatic tumours131,132. As such, formins are promising targets for cancer therapies and, indeed, T cells engineered to recognize a formin 1 (FMN1) peptide exhibit potent anti-tumour activity133.

Connections also exist between formins and immune diseases. mDia1–/– mice develop a range of myeloproliferative defects that affect T cell function130,134. This could stem from the role of mDia1 in T cell polarization towards antigen presenting cells135. FMN1 and mDia1 are also crucial for macrophage phagocytosis and immune function84,93,136. New studies are therefore needed to probe the involvement of formins in inherited immune disorders.

Formin mutations also cause defects in fertility and development. Knockout studies show that FMN2 and mDia1 are essential for normal gamete formation137–139 and embryonic development140–142, and FMN1 is essential for normal limb development143. A chromosomal truncation of human Dia2, which is homologous to mDia3 (also known as DIAPH2), is linked to premature ovarian failure144, and a distinct chromosomal truncation of human Dia1 causes non-syndromic deafness by abrogating cilia formation in hair cells145. As these examples only touch on a few of the 15 formin genes, the future is ripe for discovering new relationships between formins and disease.

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Autoinhibition

Regulatorya bActive

RBD DID

DD CC

DD CC

RBD DID

DAD

DAD

FH2

Rho andRacfamilyGTPases

α-cateninSpa2

IQGAP1

ProfilinIRSp53DIPHof1

Bud14CLIP170DIPeEF1ASmy1 Bud6

DishevelledROCK

FH1

N

NC

C

Profilin

Actin

Nature Reviews | Molecular Cell Biology

FH2

Actinfilament

F-actin(filamentous actin). A flexible, helical polymer of globular actin (G-actin) subunits, with a diameter of 5–9 nm.

Capping proteinOne of a group of ubiquitously expressed proteins that are found in eukaryotic cells and show high affinity for the barbed ends of actin filaments, thereby antagonizing filament growth.

Specialized effects. some formins have additional effects on actin beyond nucleation and elongation. For example, S. cerevisiae Bnr1, mouse mDia2, FMn-like protein 1 (FMnL1; also known as FRL1), FMnL2 (also known as FRL2) and FMnL3 (also known as FRL3) and Arabidopsis thaliana FORMIn-LIKe 1 (FH1) bundle actin fila-ments41,56–58, and mouse FMnL1 and inverted formin 2 (InF2) and A. thaliana FH8 sever and/or depolymerize actin filaments59–61. It is appealing to consider how these effects might expand the versatility of formins in vivo, but addressing this issue will probably require uncoupling these activities from nucleation and elongation effects. Bundling could be especially important in the construction of actin arrays comprised of parallel crosslinked filaments (for example, stress fibres, filopodia and cables). severing may play an important role in promoting disassembly and turnover of actin structures, an aspect of formin function that has not yet been explored in vivo.

Control of microtubule dynamics. A growing body of evidence suggests that, in addition to regulating actin dynamics, many formins directly bind and regulate microtubules. This puts formins in an ideal position to coordinate functions that depend on the close collabo-ration of the actin and microtubule cytoskeletons62,63. Members of three formin subfamilies (Dia, FMn and InF) have possible roles in microtubule regulation. It has been difficult to fully uncouple in vivo formin effects on microtubules from effects on actin, since perturbations of one system often affect the other. However, at least some effects appear to be separable (see below).

Much of our knowledge about formin regulation of microtubules has come from studies on cultured fibro blasts, which contain two separate populations of micro tubules — one stable and one highly dynamic. Microtubules have two structurally distinct ends (plus and minus ends). Dynamic microtubules grow and

Figure 2 | Domain organization of autoinhibited formins. a | The crystal structures and a schematic of formin domains, showing the proteins that bind them (with corresponding colours). The amino-terminal crystal spans the diaphanous inhibitory domain (DID) and the coiled-coil (CC) domain of mouse diaphanous 1 (mDia1; also known as DIAPH1)148. The formin homology 2 (FH2) crystal corresponds to the FH2 dimer of Bni1 (REf. 35). The carboxy-terminal crystal corresponds to the Dia autoregulatory domain (DAD) of mDia1 (REf. 22). FH2 dimerizes in an anti-parallel manner to form a doughnut-shaped structure, whereas FH1 (located between the CC and FH2 domains) lacks a predicted structure but might be rope-like. In the crystal structure of the regulatory N-terminal half, the circled areas on the DID indicate the DAD binding sites that mediate autoinhibition. Note that Rho binding to the Rho-binding domain (RBD) may relieve autoinhibitory interactions. In mouse, IQ motif-containing GTPase activating protein 1 (IQGAP1) binds the DID, insulin receptor substrate p53 (IRSp53; also known as BAIAP2) binds FH1 and cytoplasmic linker protein 170 (CLIP170; also known as CLIP1) binds FH2 of mDia1; Dia-interacting protein (DIP; also known as WISH and NCKIPSD) binds FH1 and FH2 of mDia1 and mDia2 (also known as DIAPH3); and α-catenin binds the CC domain of formin 1 (FMN1). In Saccharomyces cerevisiae, Spa2 binds the CC domain, elongation factor 1α (eEF1A) binds FH2 and bud site selection protein 6 (Bud6) binds the DAD of Bni1; and cytokinesis protein 2 (Cyk2; also known as Hof1) binds FH1 and Bud14 and Smy1 bind FH2 of Bni1-related protein 1 (Bnr1). In humans, Rho-associated protein kinase (ROCK1) binds the DAD of FH1/FH2 domain-containing protein 1 (FHOD1) and dishevelled binds the DAD of dishevelled-associated activator of morphogenesis 1 (DAAM1). Profilin binds FH1 of all formins examined. For additional details on interactions, see TABLE 1. b | Schematic of a formin dimer in action. The dimeric FH2 ‘rides’ the growing barbed end of actin through dynamic motions of its two functional halves, alternating contacts with actin subunits exposed at the filament end. The adjacent rope-like FH1 domains recruit profilin–actin complexes and deliver actin subunits to the growing barbed end (arrows). The grey curved lines emphasis the dynamic motion of the formin on the barbed end.

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ENA/VASP(Enabled/vasodilator-stimu-lated phosphoprotein). A member of a family of actin-binding proteins that are required for many cellular processes, including motility. similar to formins, ENA/VAsP proteins associate with the barbed ends of actin filaments and protect growing filaments from capping proteins, although they appear to use a distinct mechanism.

shrink primarily at their plus ends. stable microtubules are typically oriented with their plus ends facing the plasma membrane and provide secure tracks on which motor proteins traffic various cargos required for cell dynamics and motility64. Overexpression of dominant-active mDia2 or activation of endogenous mDia1 mark-edly increases the number of stabilized microtubules65. In line with a role for these formins in microtubule stabili-zation, endogenous mDia1 and mDia2 partially localize to a subset of stable microtubules at the cell periph-ery16,65, and a mDia2 construct containing FH1 and FH2 domains binds directly to and stabilizes microtubules in vitro18. Binding requires relatively high concentrations of mDia2 (6.1 μM Kd), suggesting that other factors may facilitate micro tubule association in vivo. Recently, the microtubule stabilization effects of mDia2 in vitro and in vivo were shown to be independent of FH2 domain dimerization and actin nucleation18. Importantly, these findings show that mDia2 activities on actin and micro-tubules are separable. However, the actin and microtubule functions might be closely coordinated in vivo. Although FH2-mediated actin assembly was found to be dispensa-ble for micro tubule stabilization in vivo, it was required for the proper extension of stable microtubules into the cell periphery18. This suggests that a key area for future investigation is the interplay between the formin effects on actin and microtubule dynamics.

Mechanistically, how do mDia1 and mDia2 stabilize microtubules? Based on results from injecting cells with either mDia1 or mDia2 with fluorescent tubulin, each formin appears to decrease tubulin subunit exchange at microtubule plus ends65. How this is achieved is not fully understood, but purified mDia2 alone (2 μM) is suffi-cient to retard microtubule shortening rates by approxi-mately 60% in vitro18. An equivalent biochemical analysis of mDia1 activities on microtubules is needed to better understand the mDia1 in vivo observations. Interestingly, endogenous mDia1 co-immunoprecipitates with the microtubule tip-associated proteins eB1 and APC16. Therefore, it will be interesting to learn whether APC and eB1 influence the effects of mDia1 on microtubules. Another interesting point is that, whereas purified mDia2 binds to microtubules indiscriminately along their sides in vitro18, mDia1 appears to be enriched at microtubule plus ends in vivo16. Thus, perhaps eB1 and/or APC assist in recruiting mDia1 to microtubule plus ends.

what function is served by formin-mediated stabil-ization of microtubules? In fibroblasts, mDia1 and/or mDia2 promote polarization and directed motility, possibly through interactions with microtubules16. In an interesting parallel, D. melanogaster DIA promotes microtubule stabilization at neuromuscular junctions to regulate synaptic growth66. Furthermore, the S. cerevisiae Dia-related formin, Bni1, is required for mitotic spindle

Table 2 | The seven subfamilies of mammalian formins

Subfamily* Members* N-terminal regulatory domains

Physiological functions In vitro microtubule binding

In vitro actin nucleation

Refs

DAAM DAAM1 and DAAM2 RBD and DID Widely expressed with crucial roles in vertebrate gastrulation, cellular morphogenesis and neuronal growth cone dynamics

Unknown Yes 27, 157–159

Delphilin Delphilin PDZ Neuronal-specific expression; binds to glutamate receptors at the postsynaptic side of neuromuscular junctions

Unknown Unknown 30,160

Dia mDia1 (DIAPH1), mDia2 (DIAPH3) and mDia3 (DIAPH2), and their respective homologues hDia1, hDia3 and hDia2

RBD and DID Widely expressed with roles in lammellipodial and filopodial protrusion, stress fibre formation, cell adhesion, cell migration, cytokinesis, phagocytosis, endosomal trafficking, synaptic growth and stability, microtubule stabilization and kinetochore attachment and transcriptional activation

Yes Yes 17,18,23, 25,26,34, 66,80,88,

93,99,100, 142,161

FHOD (FHOS)

FHOD1 and FHOD3 (FHOS2)

GBD and DID Widely expressed with roles in membrane protrusion, stress fibre formation, microtubule alignment with stress fibres, cell adhesion and transcriptional activation

Unknown Unknown 24,39, 162–164

FMN FMN1 and FMN2 RBD and DID Highly expressed in the developing nervous system, kidney and oocytes, with roles in membrane protrusion, cell adhesion, mitotic spindle positioning, cytokinesis and actin meshwork formation in oocytes

Yes Yes 76, 165–168

FMNL (FRL) FMNL1 (FRL1), FMNL2 (FRL2) and FMNL3 (FRL3)

GBD and DID Widely expressed with roles in phagocytosis, cell adhesion and cell migration

Unknown Yes 58,59,84, 169

INF INF1 and INF2 DID Widely expressed with roles in microtubule stabilization and organization of the endoplasmic reticulum

Yes Yes 60,74,87

DAAM, dishevelled-associated activator of morphogenesis; DID, diaphanous inhibitory domain; FHOD, FH1/FH2 domain-containing protein; FMN, formin; FMNL, FMN-like; GBD, GTPase-binding domain; hDia, human diaphanous; INF, inverted formin; mDia, mouse diaphanous; RBD, Rho-binding domain; PDZ, postsynaptic-density protein of 95 kDa, Discs large and Zona occludens 1.*Alternative protein names are provided in brackets.

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KinetochoreA large multiprotein complex that assembles onto the centromeric DNA of chromosomes and links the chromosome to microtubule plus ends in the mitotic spindle. It plays an instrumental role in chromosome segregation during mitosis.

Microtubule arrayA spatial arrangement of microtubule polymers harnessed to cellular tasks that include mitosis, intracellular transport and directional cell movement.

Amphipathic helixA helical structure that consists of hydrophobic non-polar residues on one side and hydrophilic polar residues on the other side.

orientation67, and Bud6, a ligand of Bni1, binds to micro-tubules and facilitates cortical capture of microtubule plus ends to promote nuclear migration68. In another possible parallel, mDia3 (also known as DIAPH2) is required for microtubule attachment to kinetochores and for chromo-some alignment and segregation69. Thus, multiple roles for formins in microtubule regulation are emerging. It will be important to determine which, if any, of these functions are tied to the ability of formins to stimulate actin assem-bly. Determining the localization patterns of these formins with both actin and microtubules in the same cells will help address whether formins can interact simultaneously with actin and microtubules. In principle, formins could use their microtubule plus end association to assemble polarized actin fibres that function as guide wires for myosin motors to steer microtubule ends towards cortical actin foci or kinetochores67,70–72. Formins might also directly bridge microtubule and actin polymers, aligning them in parallel arrays73–75.

Microtubule interactions have also been reported for the formins InF1 (also known as FHDC1) and D. mela-nogaster Cappuccino (CAPu)74–76. unlike other formins examined to date, endogenous InF1 localizes predomi-nantly to microtubules rather than actin structures, and overexpression of InF1 stabilizes microtubule arrays74. A specific domain in InF1, at the C-terminal side of FH2, is sufficient to induce stabilized microtubules. The micro-tubule-binding site on mDia2 maps to a similar region, which includes part of the FH2 domain plus adjacent C-terminal sequences18. In contrast to mDia2 and InF1, which stabilize microtubules, genetic evidence suggests that CAPu increases microtubule dynamics75. Purified CAPu binds to microtubules and crosslinks microtubules and actin in vitro, and is genetically implicated in regu-lating both polymers75. The microtubule-binding domain of CAPu maps to its FH2, whereas the mammalian

homologue FMn1 binds to microtubules through a domain found in its n terminus76.

In summary, microtubule-regulating formins can have distinct effects on microtubule dynamics, which may be due in part to the use of different microtubule-binding domains. In general, the in vitro affinities of formins for actin are much higher than for microtubules, but in vivo interactions with actin and microtubules could be dif-ferentially regulated by post-translational modifications and/or binding partners to promote selective inter actions. It remains to be determined whether formins bind simultaneously to microtubules and actin.

Formin regulationCells use various spatial and temporal regulatory mecha-nisms to control the effects of formins on actin assembly and microtubule dynamics. Indeed, severe defects in actin and microtubule organization result from the expression of mutated, constitutively active formins77–80. The diversity in formin binding proteins (TABLE 1) ena-bles formins to be involved in various tasks, including cell polarization, adhesion, movement and division. Below, we describe recent advances in understanding the control points in formin regulation. Little is known about how formin effects on microtubules are regulated; therefore, this discussion focuses on the regulation of formin activi-ties on actin. For clarity, we frame the discussion into a working model for the formin activity cycle (fIG. 3).

Step 1: autoinhibition. Many animal and fungal formins appear to be maintained in an inactive or ‘resting’ state in the cytosol, primarily through autoinhibition, before being recruited to and activated at membranes. Autoinhibition, which is mediated by intramolecular interactions between the formin n and C termini, has been observed in five of the seven mammalian formin subfamilies, the excep-tions being InF and delphilin. Autoinhibition is achieved through binding of the C-terminal DAD to the n-terminal DID26,78,81. Two sets of residues in the DAD have a crucial role in autoinhibition: an amphipathic helix (MDXLLXL), the hydrophobic side of which contacts a conserved sur-face on the DID26,82,83, and a nearby 3–15 residue Lys- and Arg-rich sequence, which also contributes to DID bind-ing24,25. DAD–DID interactions inhibit the ability of FH2 to nucleate actin assembly both in vitro and in vivo24,26,84. These associations are hypothesized to sterically obstruct FH2 interactions with actin, but structural validation of this model is needed.

Additional modes of regulation are possible in the FH2 domain, which we refer to here as inter-FH2 autoinhibition. The crystal structure of the FH2 of dishevelled-associated activator of morphogenesis 1 (DAAM1) shows that its two functional halves pack together in a manner that occludes actin binding85,86. Accordingly, purified DAAM1 has very weak nucleation activity, 100-fold weaker than mDia1 (REf. 85). This inter-FH2 autoinhibition in DAAM1 is maintained by a novel β-strand inter action in the FH2 linker subregion, which is absent in Bni1. Mutations designed to disrupt the β-strand interface increase DAAM1 nucleation activity, raising the possibility that specific FH2 binding partners

Box 2 | Visualization of actin filament dynamics by TIRF microscopy

The study of actin dynamics has been enriched greatly by the development of total internal reflection fluorescence (TIRF) microscopy as a tool to visualize actin dynamics at the single filament level. TIRF assays use actin monomers that are covalently labelled with fluorescent dye molecules and offer a major advantage over standard ‘bulk’ pyrene–actin assays, which measure the average behaviour of a filament population (billions per reaction). Measuring actin dynamics in bulk makes it difficult to know at any given time how many filaments in the reaction are growing versus shortening, or how many are associated with the protein being studied. The full range of effects of an actin regulator can therefore be missed. TIRF bypasses this limitation by monitoring the behaviour of individual filaments. As such, the use of TIRF has led to important breakthroughs. For instance, the ability of formins to accelerate elongation was missed in bulk assays but elegantly revealed in TIRF, in which it was possible to distinguish rapidly growing ends associated with a formin from slower growing free ends53. However, TIRF also has limitations. It requires a higher excitation level than bulk assays, which can lead to photobleaching problems. The generation of statistically relevant results by TIRF requires large data sets to be obtained, which is time consuming and computationally intense. Furthermore, in TIRF, filaments are tethered or restricted to a glass surface, which can alter polymer dynamics. For these reasons, it is ideal to combine bulk and TIRF analyses in parallel to study the effects of actin regulators.

Finally, it is worth noting that because the resolution of light microscopy is 200 nm, or 70 subunits of actin polymer, standard TIRF conditions are limited to detecting the net addition or loss of 70 subunits. Therefore, to capture crucial information about individual molecular events requires single molecule analysis.

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Nature Reviews | Molecular Cell Biology

2 Recruitment and activation

3 Nucleation 4 Elongation 5 Displacement

6 Inhibition and release

1 Autoinhibition

Actin monomer

Nucleationcofactor

Displacementfactor

DID domain

DAD domain

FH2 domain FH1 domain

Rho

Plasma membrane

Inhibitor

Profilin

Actinfilament

Rho GTPaseOne of a conserved family of small enzymes that converts GTP to GDP and acts as a ‘molecular switch’ that is active in the GTP-bound form and inactive in the GDP-bound form. Active Rho proteins bind to effectors, such as formins, to trigger cytoskeletal remodelling.

or modifications might promote release of the inter-FH2 autoinhibition to stimulate actin assembly.

There is a growing list of formins that appear to not be regulated by classic DID–DAD autoinhibition. This includes mammalian delphilin, FMnL2 and InF proteins, all plant formins, Schizosaccharomyces pombe Cdc12 and D. melanogaster CAPu6,13,30,58,75,87. Currently, there is lit-tle understanding of how the activities of these formins are controlled, but one possibility is that they are inhib-ited by specific binding partners. Indeed, three formin inhibitory ligands have been identified: Dia-interacting protein (DIP; also known as wIsH and nCKIPsD), spire and Bud14 (REfs 45,88,89). DIP is a mammalian pro-tein that directly interacts with the for mins mDia1 and mDia2, and is required for proper forma tion of cortical actin networks88. As mentioned above, D. melanogaster spire syner gizes with the formin CAPu to assemble cyto plasmic actin meshworks in oocytes44. In S. cerevi-siae, Bud14 directly interacts with the formin Bnr1 and is required for the architecture and function of actin cables89. All three of these inhibitors target FH2 and block actin assembly. However, any further relationship among their mechanisms is yet to be determined. More work is

also needed to explore the regulation of formin activities by post-translational modification.

Step 2: activation and recruitment. How formins are released from autoinhibition is not fully understood. A large body of evidence shows that Rho GTPases have cen-tral roles in activating formins. First, Rho proteins must be locally activated by signals (for example, chemoattract-ants, mating pheromones or growth factors) and then they can recruit downstream effectors, including formins, to remodel the cytoskeleton90. There are numerous differ-ent Rho proteins expressed in each eukaryotic cell type, and there is a high degree of binding specificity between particular formins and Rho proteins23,91,92. Activated Rho proteins bind to the RBD of formins and help disrupt adjacent autoinhibitory DID–DAD interactions20,22. Rho proteins are sufficient to release autoinhibited formins and promote actin assembly in vitro, but this requires high concentrations of Rho (over 8 μM), which may not be physiological26,84,93. This suggests that additional factors are involved in activating formins in vivo. Indeed, release of FMnL1 and mDia1 from autoinhibition in vivo has been shown to require unspecified membrane-associated

Figure 3 | Proposed model of regulatory points in the formin activity cycle. Formin dimers are autoinhibited in the cytosol through interactions of their amino-terminal diaphanous inhibitory domain (DID) and their carboxy-terminal diaphanous autoregulatory domain (DAD) (step 1). Formins are recruited to and activated at the plasma membrane by Rho proteins and possibly other factors (not shown). This leaves the DID associated with the plasma membrane and liberates the doughnut-shaped formin homology 2 (FH2) domain and the adjacent DAD to initiate actin assembly (step 2). Nucleation of an actin filament may involve FH2 binding and the stabilization of transient actin polymerization intermediates (step 3). In some formins, strong nucleation might require an actin monomer-binding nucleation cofactor, such as the one depicted here (based on bud site selection protein 6 (Bud6)), to associate with the DAD. The barbed end of the nascent actin filament is captured by the FH2 domain (step 4). The FH2 stays processively attached to the growing end as new actin subunits are rapidly added. Elongation is accelerated by the flanking, rope-like FH1 domains through their ability to recruit and deliver profilin–actin complexes to the growing barbed end. Interactions between the FH2 domain and a formin displacement factor trigger the release of the actin filament and its incorporation into an actin network, leaving the inhibited formin membrane-associated (step 5). The formin and its attached actin filament jointly dissociate from the membrane (step 6). This is probably coupled to formin inhibition, which is achieved through post-translational modification and/or the binding of additional cellular factors.

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FarnesylationA post-translational modification in which a farnesyl group (a hydrophobic group of three isoprene units) is conjugated to proteins, such as Ras GTPases, that contain a C-terminal CAAX motif. farnesylation promotes insertion of the modified proteins into lipid bilayers.

Phagocytic cupAn actin-rich, cup-like extension of the peripheral membrane that partially encircles foreign particles or bacteria during the early stages of the phagocytic process.

WAVE complex(WAsP-family verprolin-homologous protein complex). A complex of five proteins that regulates Arp2/3 complex-mediated actin assembly. The defining member of this complex, WAVE, is related to WAsP and directly stimulates the Arp2/3 complex to nucleate actin assembly, while other components of the complex regulate WAVE.

Adherens junctionA cell–cell adhesion complex that contains classical cadherins and catenins, which are attached to cytoplasmic actin filaments.

α-cateninA central component of adherens junctions that links cell adhesion molecules, such as cadherin, to actin filaments.

factors that cooperate with Rho84. In addition, dishev-elled (DVL) binds the DAD of DAAM1 and disrupts DID–DAD interactions either independently or with Rho27. These observations suggest that the mechanisms for activating formins are complex and involve the con-vergence of separate regulatory inputs. To clarify these mechanisms, more effort is needed to identify the full set of formin ligands, particularly those interacting with DID and DAD.

some formins appear to be released from autoinhi-bition by post-translational modification even in the absence of Rho binding. During mitotic cell growth, p53-regulating kinase 1 (Prk1), a ser/Thr protein kinase required for proper organization and function of the S. cerevisiae actin cytoskeleton, phosphorylates Bni1 at sites in its n and C termini. Phosphorylation releases Bni1 from autoinhibition for actin assembly in vitro94. In mating cells, Bni1 is phosphorylated by the mitogen-activated ser/Thr protein kinase Fus3, which regulates Bni1 localization and actin cable assembly95. Mammalian FHOD1 is phosphorylated by ROCK at three conserved ser residues near its DAD, leading to full activation of FHOD1 (REf. 39). Interestingly, ROCK also binds to FHOD1, suggesting that it may have additional roles in regulating actin nucleation (see Bud6 above).

spatially unrestricted actin assembly in cells causes severe defects in cell polarity, shape, movement and division. For this reason, formin activation must be confined to specific locations, typically at the cell cortex. In many cases, activation and localization go hand-in-hand. For example, as stated above, phosphorylation by Fus3 is required for Bni1 activation and localization95. Furthermore, in yeast and mammalian cells, Rho inter-actions regulate formin cortical localization and activa-tion42,96–99,100. what is not yet clear is whether formins are first released from autoinhibition in the cytosol and then recruited to the cortex, or whether they are locally activ ated at the cortex.

Although Rho proteins play a central part in recruit-ing formins to membranes, mounting evidence suggests that they are not always sufficient. For example, after com-plete disruption of the RBD of FMnL1 and mDia1, many formin particles were still localized at the cell cortex84. In this study, it was shown that the close dimerization and coiled-coil domains of mDia1 could help direct its locali-zation99. Other examples include the S. pombe formin Fus1, for which a DID-containing fragment was suffi-cient for localization to the cell cortex101, and S. cerevisiae Bnr1, for which domains outside of the RBD were required for localization102. Moreover, InF2 is farnesylated, which might help target it to endoplasmic reticulum membranes, possibly independently of Rho87.

which ligands of the DID, dimerization and coiled-coil domains regulate formin recruitment? In S. cerevisiae, the polarity factor spa2 helps localize Bni1 to the cortex and binds to sequences near the coiled-coil domain103. In S. pombe, the polarity factor tip elongation aberrant pro-tein 4 (Tea4) is required for the localization of formin 3 (For3) to cell tips through interactions with the RBD and flanking regions104. In mammalian cells, the RAC1- and Cdc42-effector IQ motif-containing GTPase

activating protein 1 (IQGAP1) binds to the DID of mDia1 and is required for mDia1 localization to phago-cytic cups and to the leading edge during wound healing93. Furthermore, IQGAP1 and DAD compete for DID bind-ing, suggesting that Rho and IQGAP1 may work together to activate mDia1. This could involve Rho first binding and triggering the release of DID–DAD interactions, then IQGAP1 binding to the DID to stabilize mDia1 in an active state and maintain its localization at the cor-tex. Another factor that regulates mDia1 recruitment to phagocytic cups is the microtubule plus end tracking protein CLIP170, which binds to the mDia1 FH2 (REf. 17). Interestingly, although CLIP170 recruitment of mDia1 is required for phagocytosis, CLIP170 and mDia1 colocalize only in non-phagocytosing regions of the cell. CLIP170 might be required specifically for early recruitment of mDia1 to the cortex, whereas others factors, such as IQGAP1, maintain mDia1 localization during phagocytic cup development. Thus, regulation of mDia1 localization may involve distinct temporal phases accompanied by the formation of separate complexes.

Further mechanisms target mDia1 to adhesion sites. Abelson interactor 1 (ABI1), a component of the WAVE complex, recruits mDia1 to adherens junctions105. Although direct mDia1–ABI1 binding has not been reported, the two proteins form a complex in vivo, and ABI1 recruitment of mDia1 is required for adherens junction formation105. As ABI1 also regulates wAVe–Arp2/3-mediated actin assembly, these observations point to a possible mechanism for spatially and temporally co ordinating formin- and Arp2/3-dependent activities at cell–cell junctions. Another factor that recruits formins to adhesion sites is α-catenin, which binds to the coiled-coil domain of FMn1 (REf. 106). Thus, proper formation of cell–cell adhesion sites requires the formation of multiple formin complexes.

Step 3: nucleation. Once localized and activated, formins are able to assemble actin. However, as mentioned above, the mechanism that initiates filament formation remains poorly understood and might, in some cases, involve additional regulatory inputs. For some formins the FH2 domain might be sufficient to promote strong nucleation in vivo, but in other formins nucleation might depend on the binding of actin monomers to sequences located out-side of the FH2, and/or the association of actin monomer- binding cofactors. The concept of formins depending on nucleation cofactors draws parallels with Arp2/3 regula-tion. The Arp2/3 complex has inherently poor nucleation activity and requires a cofactor — a nucleation promot-ing factor (nPF) —for efficient actin assembly107. As mentioned above, formins exhibit a range of nucleation activities (over two orders of magnitude in direct com-parisons)56,108. It is possible that in vivo nucleation by ‘weaker’ formins relies on the association with cofactors, similar to how Arp2/3 complex-mediated actin assembly relies on its actin monomer-binding cofactor wAVe. This appears to be true for Bni1, which has a nucleation activ-ity that is approximately 15-fold weaker than that of Bnr1 and mDia1 (REf. 108). Bni1-mediated actin cable assembly in vivo depends on Bud6, its actin monomer-binding

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TropomyosinOne of a conserved family of actin binding proteins that bind to the sides of actin filaments, stabilizing actin, and in some cases binds to formins to promote actin assembly.

cofactor, which increases Bni1 nucleation activity by 15-fold (A.G.D & B.L.G, unpublished observations). Live- cell micro scopy studies suggest that Bud6 and Bni1 are recruited independently to the cell cortex, where they transiently interact to assemble actin cables109,110. Thus, the co ordinated spatial regulation of a formin and its cofactor provides one effective mechanism for restricting nuclea-tion to specific locations, and is similar to how Arp2/3 and wAVe are spatiotemporally coordinated in vivo to promote localized actin assembly111.

Step 4: elongation. As described above, once an actin fila-ment has been nucleated, the formin ‘rides’ the growing barbed end, and elongation can be accelerated many fold by FH1 domain interactions with profilin–actin. Based on rates of elongation stimulated by mDia1 in vitro in the presence of profilin53, a cellular concentration of 10 μM profilin–actin would lead to mDia1 elongating filaments at 550 subunits per second in vivo — about 1.5 microns of filament per second. Indeed, this predicted rate closely matches the observed speed of actin polymerization-dependent mDia1 particle movement in living cells49.

One possible mechanism that might be used by cells to regulate elongation is competitive interactions between profilin and other proteins for binding FH1 domains. In addition to profilin-binding sites, many FH1 domains contain consensus binding sites for sH3 and ww domains. A genome wide screen for FH1 bind-ing partners of FMn1 identified a large number of sH3 and ww domain-containing proteins, named formin-binding proteins (FBPs)112. The in vivo and in vitro effects of FBPs binding to FMn1 are unclear, but are a promising avenue for future investigations. In addition, the sH3 domain-containing kinase src binds to the FH1 domains of delphilin, DAAM1, mDia2 and FMn1 (REfs

30,81,113,114). mDia2 binding activates src kinase activ-ity, but whether src binding regulates mDia2 activity is not known115. Finally, interactions have been observed between the FH1 domain of mDia1 and the sH3 domain of insulin receptor substrate p53 (IRsp53; also known as BAIAP2), a protein that coordinates membrane and actin cytoskeleton remodelling116,117. It will be important to determine the effects of IRsp53 binding on formin activity. Given the proximity and overlap of sH3 and ww domain-binding sites with profilin-binding sites in FH1 domains, there is strong potential for uncovering new regulators of formin-mediated actin elongation.

Steps 5 and 6: attenuation. The points above raise a new question — how is the duration of a formin-mediated actin assembly event regulated to control filament length? Many of the actin arrays assembled by formins in vivo consist of short filaments (0.3–2.3 microns), including cables, cytokinetic rings and stress fibres118–120. simple cal-culations suggest that these short filaments are assembled on a timescale of less than one second in vivo; however, purified formins remain bound to and elongate barbed ends for many minutes without dissociating in vitro15,53. These observations suggest that there may be active mechanisms in vivo for displacing formins from growing barbed ends, and/or inhibiting or ‘locking’ formins into

an inactive state bound to filament ends, forming a static cap. These two mechanisms are discussed below and are depicted in our model for formin regulation (fIG. 3; steps 5 and 6).

Attenuation of elongation by a displacement fac-tor was recently reported for S. cerevisiae Bud14 acting on the formin Bnr1 (REf. 89). Bnr1 molecules are stably anchored to the bud neck throughout the cell cycle109 and therefore must undergo rapid, repeated rounds of actin assembly to continuously produce the short filaments (0.3–0.5 microns) that comprise the cables that stream into the mother cell119,121. Bud14 binds strongly to the FH2 domain of Bnr1 and displaces Bnr1 from growing barbed ends in vitro, and regulates actin cable length and architecture in vivo89. However, these observations leave open the question of how Bud14–Bnr1 interactions are subsequently disrupted to recycle Bnr1 for new rounds of actin assembly and completion of the activity cycle (fIG. 3). similar displacement factors might be used in other organisms to control filament length, and might be FH2 binding partners.

In the case of stably anchored formins, such as Bnr1, displacement leaves the formin associated with the cell membrane, where it must be recycled for new rounds of actin assembly. However, other formins are dynamically recruited to membranes and released after a brief period of activity. Thus, the activities of different formins may be controlled by distinct temporal inactivation mechanisms. Two examples of ‘dynamic’ formins are S. cerevisiae Bni1 and S. pombe For3 (REfs 109,110). Live-cell imaging indi-cates that Bni1 and For3 particles are transiently recruited to cell tips and, after about 15 seconds, undergo retro-grade movement in from the cell cortex on actin cables. The formin particles move at rates similar to cable flow, suggesting that they may be inactive when associated with cables. mDia1–green fluorescent protein speckles are also dynamic and undergo rapid, actin-dependent movements along linear paths in cells49. Thus, the move-ment observed might be mDia1 particles travelling on polymerizing barbed ends.

what are the mechanisms for triggering formin release from the membrane and inhibiting formins when they are attached to cables? This is unknown, but could involve formin modifications and/or binding partners. One candidate for this role is the ligand DIP, which binds to the FH2 domain of mDia2, inhibits its actin nucleation activity in vitro and suppresses mDia2-induced filopodial extension in vivo88. Another candidate is the actin-binding protein tropomyosin, which in S. pombe was shown to promote actin filament annealing, suggesting that it might ‘trap’ formin Cdc12 molecules between annealed filaments to attenuate elongation122. Other mechanisms might exist for arresting formins on membranes and/or filament ends, involving formin binding partners that act as ‘pause’ factors.

PerspectivesIn recent years, our view of formin regulation has evolved substantially. earlier models depicted Rho proteins as single-handedly recruiting and activating formins to stimulate actin assembly and/or microtubule stabilization.

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It is now indisputable that numerous cellular factors, in addition to Rho, contribute to formin regulation, and that some formins may not be regulated by Rho at all. It has also become evident that there are many different levels of formin regulation. These include mechanisms for recruit-ing formins to distinct locations and mechanisms for con-trolling distinct points in the temporal activity cycle of formins: autoinhibition, activation, nucleation, elongation and attenuation.

Linkage to specific regulatory mechanisms is deter-mined by differences in the sequences of the n-terminal halves and DAD-containing regions of formins, and by differences in the FH1 and FH2 domains. Through these differences, formins have evolved to act down-stream of multiple signalling pathways and to be involved in a wide range of cellular tasks. This heterogeneity in

formin regulation might also enable the functions of dif-ferent formins in the same cell type to be separated or combined to increase functional versatility.

Many questions about formin mechanisms and regu-lation remain open. For example, how do formins gov-ern microtubule dynamics, and how do formin binding partners such as eB1 and APC affect these functions? In addition, a complete ‘regulatory cycle’ has not yet been described for any formin. Furthermore, it has recently emerged that the in vivo functions of some formins are controlled by protein degradation, which raises new ques-tions about the mechanism of regulated formin turnover123. Investigating these and other aspects of formin regulation should provide further insights into how cells responsively remodel their cytoskeletons to drive complex behaviours such as cell motility, morphogenesis and division.

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26. Li, F. & Higgs, H. N. The mouse Formin mDia1 is a potent actin nucleation factor regulated by autoinhibition. Curr. Biol. 13, 1335–1340 (2003).The characterization of mDia1 that provided the first biochemical demonstration of formin autoinhibition through interactions between the N‑ and C‑terminal halves of the protein.

27. Liu, W. et al. Mechanism of activation of the Formin protein Daam1. Proc. Natl Acad. Sci. USA 105, 210–215 (2008).

28. Habas, R., Kato, Y. & He, X. Wnt/Frizzled activation of Rho regulates vertebrate gastrulation and requires a novel Formin homology protein Daam1. Cell 107, 843–854 (2001).

29. Kitzing, T. M. et al. Positive feedback between Dia1, LARG, and RhoA regulates cell morphology and invasion. Genes Dev. 21, 1478–1483 (2007).

30. Miyagi, Y. et al. Delphilin: a novel PDZ and formin homology domain-containing protein that synaptically colocalizes and interacts with glutamate receptor δ2 subunit. J. Neurosci. 22, 803–814 (2002).

31. Matsuda, K., Matsuda, S., Gladding, C. M. & Yuzaki, M. Characterization of the δ2 glutamate receptor-binding protein delphilin: splicing variants with differential palmitoylation and an additional PDZ domain. J. Biol. Chem. 281, 25577–25587 (2006).

32. Deeks, M. J. et al. Arabidopsis group Ie formins localize to specific cell membrane domains, interact with actin-binding proteins and cause defects in cell expansion upon aberrant expression. New Phytol. 168, 529–540 (2005).

33. Pring, M., Evangelista, M., Boone, C., Yang, C. & Zigmond, S. H. Mechanism of formin-induced nucleation of actin filaments. Biochemistry 42, 486–496 (2003).

34. Moseley, J. B. et al. A conserved mechanism for Bni1- and mDia1-induced actin assembly and dual regulation of Bni1 by Bud6 and profilin. Mol. Biol. Cell 15, 896–907 (2004).Characterization of yeast Bni1 and mouse Dia1 formins, showing that formins have a conserved function in protecting the growing barbed ends of actin filaments from capping proteins, that the FH2 domain must dimerize to be active and that Bud6 binds directly to Bni1 to stimulate its actin assembly activity.

35. Xu, Y. et al. Crystal structures of a formin homology-2 domain reveal a tethered dimer architecture. Cell 116, 711–723 (2004).This study provided the first crystal structure of an active formin molecule, revealing that the FH2 is a flexibly tethered dimer, which led to the ‘stair‑stepping’ model for processive motion of the FH2 on growing barbed ends of actin filaments.

36. Otomo, T. et al. Structural basis of actin filament nucleation and processive capping by a formin homology 2 domain. Nature 433, 488–494 (2005).

37. Sept, D. & McCammon, J. A. Thermodynamics and kinetics of actin filament nucleation. Biophys. J. 81, 667–674 (2001).

38. Higashida, C. et al. G-actin regulates rapid induction of actin nucleation by mDia1 to restore cellular actin polymers. J. Cell Sci. 121, 3403–3412 (2008).

39. Takeya, R., Taniguchi, K., Narumiya, S. & Sumimoto, H. The mammalian formin FHOD1 is activated through phosphorylation by ROCK and mediates thrombin-induced stress fibre formation in endothelial cells. EMBO J. 27, 618–628 (2008).Using complementary in vitro and in vivo approaches, this study showed that phosphorylation of the formin FHOD1 by ROCK disrupts FHOD1 autoinhibition independently of Rho binding and stimulates stress fibre formation.

40. Hannemann, S. et al. The Diaphanous-related formin FHOD1 associates with ROCK1 and promotes Src-dependent plasma membrane blebbing. J. Biol. Chem. 283, 27891–27903 (2008).

41. Moseley, J. B. & Goode, B. L. Differential activities and regulation of Saccharomyces cerevisiae formin proteins Bni1 and Bnr1 by Bud6. J. Biol. Chem. 280, 28023–28033 (2005).

42. Martin, S. G., Rincon, S. A., Basu, R., Perez, P. & Chang, F. Regulation of the formin for3p by cdc42p and bud6p. Mol. Biol. Cell 18, 4155–4167 (2007).

43. Pechlivanis, M., Samol, A. & Kerkhoff, E. Identification of a short Spir interaction sequence at the C-terminal end of formin subgroup proteins. J. Biol. Chem. 284, 25324–25333 (2009).

44. Dahlgaard, K., Raposo, A. A., Niccoli, T. & St. Johnston, D. Capu and Spire assemble a cytoplasmic actin mesh that maintains microtubule organization in the Drosophila oocyte. Dev. Cell 13, 539–553 (2007).

45. Quinlan, M. E., Hilgert, S., Bedrossian, A., Mullins, R. D. & Kerkhoff, E. Regulatory interactions between two actin nucleators, Spire and Cappuccino. J. Cell Biol. 179, 117–128 (2007).

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46. Pruyne, D. et al. Role of formins in actin assembly: nucleation and barbed-end association. Science 297, 612–615 (2002).Together with reference 14, this study was the first to show that formins directly nucleate actin polymerization, and showed unexpectedly that formins physically associate with the dynamic barbed ends of actin filaments.

47. Zigmond, S. H. et al. Formin leaky cap allows elongation in the presence of tight capping proteins. Curr. Biol. 13, 1820–1823 (2003).

48. Kovar, D. R. & Pollard, T. D. Insertional assembly of actin filament barbed ends in association with formins produces piconewton forces. Proc. Natl Acad. Sci. USA 101, 14725–14730 (2004).This study used TIRF microscopy to visualize in real time polymerizing actin filaments associated with formins at their barbed ends, and found that formins remain persistently attached to the barbed ends while allowing the insertional assembly of new subunits.

49. Higashida, C. et al. Actin polymerization-driven molecular movement of mDia1 in living cells. Science 303, 2007–2010 (2004).This study used live‑cell imaging to track the movements of mDia1–GFP fusion proteins in cultured cells, which revealed rapid and persistent, actin‑dependent movements on linear tracks, providing the first in vivo evidence for formins moving processively on growing barbed ends.

50. Vidali, L. et al. Rapid formin-mediated actin-filament elongation is essential for polarized plant cell growth. Proc. Natl Acad. Sci. USA 106, 13341–13346 (2009).

51. Breitsprecher, D. et al. Clustering of VASP actively drives processive, WH2 domain-mediated actin filament elongation. EMBO J. 27, 2943–2954 (2008).

52. Romero, S. et al. Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis. Cell 119, 419–429 (2004).This study was the first to report that formins can accelerate elongation at barbed ends, a conclusion that was reached by comparing the lengths of actin filaments assembled in the presence and absence of formins.

53. Kovar, D. R., Harris, E. S., Mahaffy, R., Higgs, H. N. & Pollard, T. D. Control of the assembly of ATP- and ADP-actin by formins and profilin. Cell 124, 423–435 (2006).This study used TIRF microscopy to show that formins markedly accelerate barbed end elongation through physical interactions between their FH1 domains and profilin–actin complexes, and that different formins support different elongation rates.

54. Vavylonis, D., Kovar, D. R., O’Shaughnessy, B. & Pollard, T. D. Model of formin-associated actin filament elongation. Mol. Cell 21, 455–466 (2006).

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57. Michelot, A. et al. The formin homology 1 domain modulates the actin nucleation and bundling activity of Arabidopsis FORMIN1. Plant Cell 17, 2296–2313 (2005).

58. Vaillant, D. C. et al. Interaction of the N- and C-terminal autoregulatory domains of FRL2 does not inhibit FRL2 activity. J. Biol. Chem. 283, 33750–33762 (2008).

59. Harris, E. S., Li, F. & Higgs, H. N. The mouse formin, FRLα, slows actin filament barbed end elongation, competes with capping protein, accelerates polymerization from monomers, and severs filaments. J. Biol. Chem. 279, 20076–20087 (2004).

60. Chhabra, E. S. & Higgs, H. N. INF2 is a WASP homology 2 motif-containing formin that severs actin filaments and accelerates both polymerization and depolymerization. J. Biol. Chem. 281, 26754–26767 (2006).

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65. Palazzo, A. F., Cook, T. A., Alberts, A. S. & Gundersen, G. G. mDia mediates Rho-regulated formation and orientation of stable microtubules. Nature Cell Biol. 3, 723–729 (2001).This study was the first to show that formins can bind directly to microtubules and regulate microtubule stability in vivo.

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AcknowledgementsWe are grateful to J. Moseley, I. Sagot and members of the Goode laboratory for helpful discussions and editing the manuscript, and A. Alberts for sharing information before publication. We apologize to colleagues whose work was not highlighted owing to space limitations. This work was sup-ported in part by grants from the National Institutes of Health to B.G. (GM63691 and GM083137).

Competing interests statement.The authors declare no competing financial interests.

DATABASESUniProtKB: http://www.uniprot.orgABI1 | APC | Bni1 | Bnr1 | Bud6 | Bud14 | CAPU | Cdc12 | CLIP170 | COBL | DAAM1 | DIA | DIP | EB1 | FH1 | FH8 | FHOD1 | FMN1 | FMN2 | FMNL1 | FMNL2 | FMNL3 | For3 | Fus1 | Fus3 | INF2 | IQGAP1 | IRSp53 | JMY | mDia1 | mDia2 | mDia3 | Prk1 | Spa2 | SPIRE1 | Tea4

FURTHER INFORMATIONBruce L. Goode’s homepage: http://www.bio.brandeis.edu/goodelab

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