Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Mol Cells. Author manuscript; available in PMC 2014 February 3.
Published in final edited form as:
PMCID: PMC3910092

Regulation of Actin Cytoskeleton Dynamics in Cells


The dynamic remolding of the actin cytoskeleton is a critical part of most cellular activities, and malfunction of cytoskeletal proteins results in various human diseases. The transition between two forms of actin, monomeric or G-actin and filamentous or F-actin, is tightly regulated in time and space by a large number of signaling, scaffolding and actin-binding proteins (ABPs). New ABPs are constantly being discovered in the post-genomic era. Most of these proteins are modular, integrating actin binding, protein-protein interaction, membrane-binding, and signaling domains. In response to extracellular signals, often mediated by Rho family GTPases, ABPs control different steps of actin cytoskeleton assembly, including filament nucleation, elongation, severing, capping, and depolymerization. This review summarizes structure-function relationships among ABPs in the regulation of actin cytoskeleton assembly.

Keywords: actin-binding protein, actin cytoskeleton, BAR protein, F-actin, filament cross-linking, filament nucleation and elongation, G-actin, Rho-GTPase, WH2


Cell movement is a vital phenomenon in most biological processes, including embryonic morphogenesis, immune surveillance, angiogenesis and tissue repair and regeneration (Hussey et al., 2006; Itoh and Yumura, 2007; McMahon and Gallop, 2005; Puppo et al., 2008; Yamaguchi and Condeelis, 2007). Actin cytoskeleton dynamics plays a crucial part in most of these processes, mediating the formation of cellular structures such as lamellipodia, filopodia, stress fibers and focal adhesions (Bailly and Condeelis, 2002). A characteristic feature of all these processes is the dynamic transition of the cellular actin between its monomeric (G-actin) and filamentous (F-actin) forms (Fig. 1C). Actin is an ATPase and ATP hydrolysis by actin plays an essential role in regulating this transition (Fig. 1A). Thus, the actin filament is asymmetric; actin monomers join the barbed (or +) fast growing end of the filament in the ATP-bound state and depart the filament preferentially from the pointed (or −) end primarily in the ADP state, giving rise to a process known as actin filament threadmilling (Figs. 1B and 1C). In addition, the transition between G- and F-actin is tightly regulated in cells by a large number of G- and F-Actin-Binding Proteins (ABPs) (Fig. 1C).

Fig. 1
Monomeric and filamentous actin. (A) The actin monomer consists of two major and structurally related domains, which because of their relative positions within the actin filament are known as the outer and inner domains. The two major domains are in turn ...

ABPs carry out a wide range of functions, including actin filament nucleation, elongation, severing, capping, and crosslinking and actin monomer sequestration (Fig. 2) (Pollard and Borisy, 2003). The reorganization of the actin cytoskeleton is regulated in time and space by multiple factor, most notably Rho family GTPases that act as GTP-dependent molecular switches (Raftopoulou and Hall, 2004). Among the small GTPases of the Rho family, Cdc42, Rac, and Rho are recognized as the most important regulators of actin assembly, controlling respectively the formation of filopodia, lamellipodia, and stress fibers (Etienne-Manneville and Hall, 2002). Signals transmitted through these GTPases lead to localized actin cytoskeleton assembly/disassembly at the plasma membrane, with the actin filaments acting to push the cellular membrane (Fig. 2) (Hall, 1994). Generally, ABPs are modular polypeptides, which in response to signaling cues undergo conformational changes, and further transmit these signals to downstream cytoskeletal partners and membranes. This is the case for example with the well-studied open/closed transition of the Arp2/3 complex Nucleation Promoting Factor (NPF) protein WASP (Figs. 3 and and5)5) (Takenawa and Suetsugu, 2007). The general outcome from these mechanisms is rapid changes in actin polymerization/depolymerization near the plasma membrane. Therefore, proteins such as the Bin/Amphiphysin/Rvs (BAR) family that link the cytoskeleton to signaling and membranes have lately attracted significant attention (Fig. 7) (Dawson et al., 2006; Frost et al., 2007; Takenawa and Suetsugu, 2007).

Fig. 2
Different actin filament networks in cells. Rho-family GTPases, including Rho, Rac, and Cdc42, are master regulators of the actin cytoskeleton. Ligand-stimulated transmembrane receptors activate Rho-family GTPases, whose hydrolysis cycle is modulated ...
Fig. 3
Internal auto-inhibition and activation by Rho-family GTPases are common themes among cytoskeletal proteins. Internal auto-inhibitory interactions in WASP (A), formin (B), and IRSp53 (C) are counteracted by Rho-family GTPases. In the case of WASP, Cdc42-mediated ...
Fig. 5
Proteins involved in actin filament nucleation and elongation. (A) Model of activated Arp2/3 complex, with bound N-WASP WCA and actin, derived from Small Angle X-ray Scattering [adapted from Boczkowska et al. Structure. 2008]. By itself, Arp2/3 complex ...
Fig. 7
BAR domain-containing proteins. (A) The BAR domain consists of an elongated anti-parallel dimer of two helical bundles, with a curved, positively charged membrane-binding interface. However, there is significant variation within this fold, and three subclasses ...

With so many cellular functions depending on the actin cytoskeleton, it is not surprising that abnormal regulation or functioning of cytoskeletal components is often a cause (or a cofactor) of numerous diseases, including cancer, neurological disorders, cardiomyopathies, cholangiocyte, glomerulosclerosis, Wiskott-Aldrich syndrome (Condeelis et al., 2005; Doctor and Fouassier, 2002; Huang et al., 2008; Machesky and Insall, 1998; Myers et al., 2006; Yamaguchi and Condeelis, 2007). While significant knowledge of the mechanisms controlling actin cytoskeleton dynamics has accumulated during the past decade from cellular and biochemical studies, detailed structural information is often lacking. Here, we summarize recent progress in our understanding of the structure-function of ABPs.

Monomeric and filamentous actin

In cells, actin exists in two states, the monomeric and filamentous states. Actin cytoskeleton dynamics is regulated by controlling the homeostatic balance between these two forms of actins, in response to extracellular stimuli (Fig. 1) (Ridley and Hall, 1992; Ridley et al., 1992). Eukaryotic actin is highly conserved in evolution from yeast to humans. Its atomic structure consists of two major domains, each consisting of two smaller subdomains (Graceffa and Dominguez, 2003; Holmes et al., 1990; Otterbein et al., 2001). According to their arrangement within the actin filament, the two major domains are known as the outer (comprising subdomains 1 and 2) and inner (comprising subdomains 3 and 4) domains (Fig. 1A). The two major domains are structurally related and might have emerged from a gene duplication event early in evolution.

The actin filament is asymmetric, and can be described as either a single left-handed short-pitch helix, with consecutive lateral subunits staggered with respect to one another by half a monomer length, or two right-handed long-pitch helices of head-to-tail bound actin subunits. Actin is an ATPase and nucleotide hydrolysis by actin helps regulate the transition between its G- and F-actin states in a process known as actin filament threadmilling (Fig. 1C). At steady state and a given Gactin concentration, F-actin grows at the barbed (+) end by spontaneous addition of ATP-bound G-actin and shortens (or depolymerizes) at the pointed (−) end by dissociation of (primarily) ADP-bound G-actin (Pollard et al., 2000).

The asymmetry of the actin filament (or directionality of growth) depends on the polarity of the actin monomers controlled by nucleotide-dependent conformational changes, which induce moderate but accumulatively important changes in the structure and stability of the filament (Belmont et al., 1999; Egelman, 1994; Scoville et al., 2006). Thus, for instance, the DNase I-binding loop (or D-loop) within subdomain 2 plays a critical role in monomer-monomer contacts within the filament. Changes in subdomain 2 and the D-loop occurring during ATP hydrolysis are thought to change the stability of actin monomers in the filament, thus promoting their dissociation (Figs. 1A and 1B) (Egelman, 1994; Holmes et al., 1990; Oda et al., 2009; Otterbein et al., 2001; Schutt et al., 1993; Scoville et al., 2006). However, a high-resolution structure of the actin filament is not yet available, and thus the exact nature of the transition is not fully understood. Since actin has a natural tendency to self-associate under physiological salt conditions, obtaining a homogeneous distribution of actin filaments for crystallization has proven extremely difficult. A potential solution to this problem is being pursued by various laboratories, by using ABPs to build ‘rulers’ to control the length distribution of actin filaments, and thereby produce monodisperse species for structural analysis.

Rho-family GTPase and the regulation of actin-cytoskeleton dynamics

During the past decade, a key question in the cytoskeleton field has been the identification of specific factors triggering cytoskeleton rearrangement in time and space. Although the molecular mechanisms remain to be fully elucidated, it is now generally accepted that signals received by cell surface receptors via chemical messengers such as cytokines, growth factors, and hormones, are transmitted to Rho-family GTPases, in particular Rho, Rac, and Cdc42 (Fig. 2) (Foster et al., 1996; Hall, 1994). The information from specifically integrated signaling pathways is then dispatched to cytoskeleton effector proteins, resulting in a myriad of processes such as acto-myosin movement (Ridley and Hall, 1992), lamellipodia (Ridley et al., 1992) and filopodia formation (Kozma et al., 1995). Consistent with this model, the ectopic expression of dominant negative mutants of Rho-family GTPases in mammalian cells severely impairs actin cytoskeleton dependent processes, including cell migration (Nobes and Hall, 1999), cytokinesis (Mabuchi et al., 1993; Prokopenko et al., 2000), endo/exocytosis (Caron and Hall, 1998), axon guidance (Luo et al., 1997), and morphogenesis during development (Lu and Settleman, 1999; Settleman, 1999). The presence of Rho-family GTPases and their effector proteins at specific loci on the membrane dictates the directionality of cell migration, with the barbed ends of the actin filaments oriented toward the membrane.

Significant effort has been devoted to identifying the specific cytoskeleton effectors of Rho family GTPases. It is well known that a large number of proteins interacting with Cdc42 and Rac contain a short stretch of ~18 amino acids referred to as the Cdc42/Rac Interactive Binding (CRIB) motif (Burbelo et al., 1995). Database searches for CRIB motif-containing proteins have identified a number of potential Rho-family effector proteins, including WASP, formins, and IRSp53 (Burbelo et al., 1995; Yamagishi et al., 2004; Zigmond, 2004b). These proteins are typically modular, i.e. they contain multiple domains, including actin-binding domains such as the WH2 and FH2 domains, and protein-protein interaction or scaffolding modules (Fig. 3). Another common trait is that these proteins are typically self-inhibited by internal interactions, which are commonly released by the binding of the GTPases. Thus, the binding of Rho GTPases accomplishes two major functions, activation and recruitment to specific loci at the membrane.

A well-studied example is the activation of the Apr2/3 complex NPF protein WASP by the Rho GTPase Cdc42 (Takenawa and Suetsugu, 2007) (Fig. 3A). In the resting state, the C-terminal WCA domains of WASP is masked by intramolecular interaction with the CRIB motif located toward the N-terminus of WASP. This conformation is referred to as the closed or auto-inhibited conformation. Upon stimulation, WASP is recruited to the plasma membrane by binding simultaneously to GTP-Cdc42, through its CRIB domain, and to the plasma membrane, through its Basic (positively charged) domain located near the CRIB domain (Prehoda et al., 2000). The activated conformation of WASP, known as the open conformation, releases the WCA inhibition, allowing for the recruitment and activation of polymerization through the Arp2/3 complex.

Formins are thought to be activated in a similar manner (Fig. 3B). Thus, in formins of the mammalian diaphanous (mDia) family, the autoinhibited conformation results from internal interaction between the dimerization domain (DID), located near the N-terminus, and the diaphanous autoregulatory domain (DAD), located near the C-terminus. The binding of GTP-Rho to the G-protein-Binding Domain (GBD), located immediately N-terminal to the DID, releases this inhibitory interaction, freeing the polymerization activity of the formin (Eisenmann et al., 2007; Goode and Eck, 2007; Peng et al., 2003; Watanabe et al., 1997).

Other than Rho GTPases, phosphorylation/dephosphorylation frequently regulates actin cytoskeleton assembly. A well-studied example is cofilin phosphorylation, which abolishes its actin-binding activity, thereby reducing filament breakdown. Ultimately, however, cofilin is also regulated by Rho GTPases, albeit in a less direct way. Thus, Rho regulates cofilin activity via Rho kinase (ROCK) that phosphorylates LIM kinase (LIMK), which in turn phosphorylates cofilin (Fig. 2) (Arber et al., 1998; Bamburg, 1999; Edwards et al., 1999; Maekawa et al., 1999). This signal transduction pathway modulates actin assembly in many cell types in response to various extracellular stimuli (Yang et al., 1998; Scott, 2007 #214).

Actin-binding proteins (ABPs)

Dynamic actin threadmilling near the cell membrane is an essential part of cell motility. This process can be subdivided into separate events, including filament nucleation, elongation, depolymerization, capping, severing, crosslinking, and actin monomer sequestration (Fig. 2) (Pollard and Borisy, 2003; Rafelski and Theriot, 2004; Zigmond, 2004a). Each of these events is controlled by specific subsets of ABPs. Structural analysis suggests that ABPs belong to a limited number of folds, including the actin-depolymerizing- factor/cofilin (ADF/cofilin) (Lappalainen et al., 1998), Wiskott-Aldrich syndrome protein (WASP)-homology domain 2 (WH2) (Paunola et al., 2002), gelsolin-homology domain (McGough et al., 2003), calponin-homology (CH) domain (Gimona et al., 2002), formin homology 2 domain (FH) (Goode and Eck, 2007). These proteins can bind G-, F-actin, or both. A special group of ABPs are the members of the myosin superfamily (Sellers, 2000), which are molecular motors that use F-actin as a track for motility. Many other proteins bind F-actin, and appear to use the actin filaments as a scaffold for their activities.

G-actin-binding proteins

Proteins that bind the actin monomer play a critical role in controlling the pool of unpolymerized actin in cells, by sequestering actin monomers and thereby modulating the pointed and barbed end addition/dissociation of actin monomers. Representative members of this group include the WH2-related protein Thymosin-beta-4 (Tβ4), profilin and ADF/cofilin (Fig. 4). The structures of various complexes of actin with ABPs have been determined (Chereau et al., 2005; Domanski et al., 2004; Hertzog et al., 2004; McLaughlin et al., 1993; Paavilainen et al., 2008; Schutt et al., 1993). A common feature among most of these proteins is the presence of a structurally conserved amphipathic α-helix that binds in the hydrophobic, or target-binding, cleft located between subdomains 1 and 3 at the barbed end of the actin monomer (Fig. 4) (Dominguez, 2004; 2007).

Fig. 4
ABPs frequently present α-helix that binds in the hydrophobic cleft of the actin monomer. (A) Structure-based alignment of the sequences of the helices of ABPs that bind in the hydrophobic cleft of actin. Hydrophobic residues directly implicated ...

The WH2 domain from diverse proteins, including ciboulot, missing-in-metastasis (MIM), and WASP-family proteins, display a similar architecture consisting of an N-terminal α-helix followed by a extended region featuring the “LKKT(V)” signature sequence motif (Fig. 4A) (Chereau et al., 2005; Dominguez, 2007; Hertzog et al., 2004; Lee et al., 2007). The hydrophobic side of the N-terminal α-helix faces the hydrophobic cleft in the actin monomer. Despite significant length and sequence variability, the C-terminal portions of the WH2 domains of different proteins follow a similar path on the actin monomer, streaming toward the pointed end of the actin monomer, i.e. towards subdomains 2 and 4. The interactions of the LKKT(V) motif and the C-terminal extended region of the WH2 domain are for the most part electrostatic in character, with positively charged amino acids of the WH2 domain facing negatively charged amino acids on the actin surface (Fig. 4B).

The structures of actin complexes with the filament capping and severing protein gelsolin (McLaughlin et al., 1993) and the monomer sequestering protein vitamin D-binding protein (DBP) (Otterbein et al., 2002) also show a similar interaction, i.e. both proteins present helices that bind in the hydrophobic cleft of the actin monomer (Dominguez, 2004). Gelsolin helix S70 to N89 and DBP helix S194 to D204 contain hydrophobic residues that bind in the hydrophobic cleft between actin subdomains 1 and 3 (Otterbein et al., 2002). Although the contact interface with actin is much larger for DBP that gelsolin, this helix has been proposed to be the most critical element of their respective interactions with actin (Figs. 4A and 4C). Furthermore, it has been suggested that the existence of this overlapping interaction in two structurally and functionally unrelated proteins allows DBP to sequester actin by forcing gelsolin out of its complex with actin, thereby allowing DBP to proceed with its actin “scavenger” function in the bloodstream (Otterbein et al., 2002). Indeed, owing to its natural tendency to polymerize, the increased concentration of actin monomers in the bloodstream during tissue injury can have devastating effects, and DBP is thought to help remove excess actin from the bloodstream by accelerating its degradation (Janmey et al., 1986).

ADF/cofilin is structurally related to gelsolin (Fedorov et al., 1997; McLaughlin et al., 1993). Early structural and biochemical studies had indicated that members of the ADF/cofilin family, including ADF1 (Bowman et al., 2000), cofilin (Fedorov et al., 1997), and twinfilin (Paavilainen et al., 2002) (Fig. 3A), also contained a helix that bound in the hydrophobic cleft of actin (Dominguez, 2004). This prediction was recently confirmed by the determination of the crystal structure of a complex of actin with twinfilin’s C-terminal ADF domain (Figs. 4A and 4C) (Paavilainen et al., 2008), as protein consiting of two ADF/cofilin domains, which binds ADP-actin monomers and filament barbed-end (Helfer et al., 2006; Ojala et al., 2002). In the structure, the helix comprising twinfilin residues I266 to S274 binds in the hydrophobic cleft of actin. Twinfilin residues Q176, at the beginning of the ADF domain, and S274 are highly conserved in the ADF/cofilin family and play critical roles in the interaction with filamentous actin (Grintsevich et al., 2008; Guan et al., 2002; Lappalainen et al., 1997), suggesting that the structure also provides a reasonable model for the interaction of ADF/ cofilin with the actin filament (Figs. 3A and and4C4C).

As illustrated by the examples above, despite the lack of overall sequence similarity among ABPs, the hydrophobic cleft in actin is emerging as the most important determinant of their interactions with actin, which is probably due to the following reasons. First, the hydrophobic cleft on actin displays remarkable plasticity; regardless of the specific amino acid sequence and directionality of binding (inwards or outwards), the helices of the various ABPs discussed above superimpose well in the hydrophobic cleft of actin (Fig. 4B). Thus, for instance, the helices of WH2 domains and ciboulot (Chereau et al., 2005; Dominguez, 2004; Hertzog et al., 2004) run in opposite direction (back to front according to the classical view) to those of gelsolin (McGough et al., 2003) and DBP (front to back) (Otterbein et al., 2002), but their main backbones overlap well. Second, actin could be capable of accommodating simultaneously interactions with functionally distinct ABPs. The relatively long helix (four helix turns) of ciboulot spans almost the entire region of the cleft (Hertzog et al., 2004), such that its N-terminal end would partially overlap with profilin, which binds at the back of the cleft (Schutt et al., 1993). In contrast, the shorter helices of gelsolin and most WH2 domains occupy only the front half of the cleft, indicating that the hydrophobic cleft could potentially accept two ABPs simultaneously in vivo. Such a co-binding mechanism has been demonstrated at least in vitro for profilin and Tβ4 (Fig. 4C) (Yarmola et al., 2007). The common actin-binding pocket also suggests a mechanism of regulation by competition, whereby ABPs compete for binding to the target-binding cleft downstream of signaling pathways, as proposed for the myocardin-related transcription factor (MRTF) (Vartiainen et al., 2007).

The Arp2/3 complex and filament nucleation

Actin filament nucleation and elongation factors that regulate the de novo formation of actin filaments in cells have received significant attention lately. Since the identification of the Arp2/3 complex (Machesky et al., 1994), a series of nucleators have been discovered, including formins (Pruyne et al., 2002), Spire (Quinlan et al., 2005), Cobl (Ahuja et al., 2007), VopL (Liverman et al., 2007), VopF (Tam et al., 2007), and Lmod (Chereau et al., 2008), as well as the elongation factors Ena/VASP (Machner et al., 2001). With the exception of formins (Fig. 5C), these molecules all use the WH2 domain for interaction with actin, which in some cases, including Spire, Cobl and VopL/VopF, takes the form of tandem WH2 repeats. In this section, we discuss structural insights into the mechanisms of nucleation of NPFs-Arp2/3 complex. For a more in-depth discussion of other WH2-based filament nucleators the reader can refer to other reviews (Dominguez, 2009; Renault et al., 2008).

In cells, nucleation, i.e. the formation of small oligomers of two to four actin subunits, is kinetically unfavorable. Moreover, actin monomers are frequently associated with actin-monomer-binding proteins such as profilin and Tβ4, which control their incorporation into filaments. Together, these two factors limit the spontaneous nucleation of actin filaments in cells, thereby creating an opportunity for cells to actively regulate the de novo polymerization of actin by using actin filament nucleation and elongation factors. The Arp2/3 complex, which consists of seven proteins including the actin-related proteins Arp2, Arp3 and subunits ARPC1-5, is the most extensively studied of the filament nucleators (Fig. 5A). By itself, the Arp2/3 complex displays very low nucleation activity (Mullins et al., 1998), but it is activated by members of a large family of Nucleation Promoting Factors (NPFs). Classical NPFs, such as the proteins WASP/WAVE, contain the C-terminal motif WCA (Machesky and Gould, 1999; Machesky et al., 1999). This region consists of three distinct segments: W, C and A. W binds the first actin subunit of the new filament (Figs. 3A and and5A).5A). The C (central or connecting) and A (acidic) motifs interact with various subunits of the Arp2/3 complex, helping to stabilize the activated conformation. The actin monomer bound to the W domain, together with Arp2 and Arp3, are thought to form a trimeric seed for the nucleation of a filament branch that emerges at a 70° angle from the side of a preexisting (mother) filament (Fig. 5A). Thus, in addition to recruiting the first actin subunits (between one and three actin subunits depending on the number of W domains of each specific NPF protein that varies between one and three), NPFs are needed to promote a conformational change within the Arp2/3 complex itself. Indeed, in the inactive structure of the Arp2/3 complex (Robinson et al., 2001), the Arps are separated and their nucleotide-binding clefts are wide open. It is believed that during activation the Arps adopt an actin filament-like conformation. In other words, according to this model (Robinson et al., 2001), Arp2 and Arp3 are the first two subunits at the pointed end of the new filament branch.

Ena/VASP and filament elongation

Ena/VASP family of proteins are WH2-based filament elongation factors, implicated in multiple cellular functions such as axon guidance and the migration of cancer cells (Brindle et al., 1996). These proteins form tetramers through their C-terminal coiled coil (C-C) domain (Bachmann et al., 1999; Kuhnel et al., 2004), and are thought to bind to the barbed ends of actin filament bundles (multiple parallel filaments), catalyzing their synchronized elongation against the plasma membrane (Fig. 2) (Brindle et al., 1996; Krause et al., 2003; 2004). In this way, the filament bundles grow to form cellular structures such as filopodia, which are actin-rich finger-like projections used by cells to sense the environment (Mattila and Lappalainen, 2008).

Like most cytoskeletal proteins, Ena/VASP proteins are modular (Fig. 5B), containing N-terminal Ena/VASP Homology 1 (EVH1), central Pro-rich and C-terminal EVH2 domains (Ferron et al., 2007). The EVH1 domain has a Pleckstrin Homology (PH)-like fold and binds the consensus sequence motif FPPPP in target proteins, helping to localize Ena/VASP proteins to their sites of action. The EVH2 region can be subdivided into G- and F-actin Binding (GAB and FAB) domains, which are both WH2-related sequences, and the C-terminal coiled coil (C-C) tetramerization domain. The central Pro-rich region binds signaling/regulatory proteins and profilin-actin. Such Pro-rich sequences are frequently found among cytoskeletal proteins, including formins and the NPFs of the Arp2/3 complex. The recruitment of multiple profilin-actin complexes to Pro-rich sequences may contribute to increasing the local concentration of actin monomers for polymerization (Kovar et al., 2006).

Structural and biochemical data suggest a molecular model of processive filament elongation by Eva/VASP proteins (Breitsprecher et al., 2008; Chereau and Dominguez, 2006; Ferron et al., 2007; Pasic et al., 2008). According to this model, profilin-actin complexes, the predominant form of polymerization competent actin in cells, are recruited to the Pro-rich region of Ena/VASP and then transferred to the adjacent GAB domain (Fig. 5B). From the GAB domain, the actin monomers can then join the barbed ends of the actin filaments tethered by the FAB domains of the Ena/VASP tetramer. With each monomer addition to the tethered filament barbed ends, Ena/VASP proteins would be expected to step forward (toward the membrane). This is probably accomplished by a mechanism of selective binding, depending on affinity modulation of the GAB and FAB domains respectively to actin monomers and the barbed ends of the actin filaments. Tetramerization is also critical for Ena/VASP function (Bachmann et al., 1999; Kuhnel et al., 2004), presumably by allowing each arm of the Ena/VASP tetramer to release and step forward while the other arms remain bound to the bundle (Ferron et al., 2007). In this way, the Ena/VASP tetramer can processively “track” the barbed end of the actin bundle (Breitsprecher et al., 2008).


Among filament nucleation/elongation factors, formins mediate the assembly of unbranched actin networks, such as filopodia and stress fibers. The diaphanous-related formins (DRFs), including mDia and Bni1p, are the best studied. Like most cytoskeletal proteins, DRFs are multidomain, multifunctional proteins, comprising GTPase-binding domain (GBD), diaphanous inhibitory domain (DID), dimerization domain (DD), coiled coil (CC), formin homology (FH) 1 and 2 domains, and diaphanous autoregulatory domain (DAD) (Figs. 3B and and5C).5C). These domains play dedicated roles in regulation, dimerization, actin nucleation/elongation and auto-inhibition (Castrillon and Wasserman, 1994; Zigmond, 2004b). In the resting state, formins exist in a folded autoinhibited conformation stabilized by internal DAD-DID interaction (Fig. 3B). The FH2 dimer, responsible for nucleation and elongation, consists of two rod-shaped domains connected in head-to-tail fashion by flexible linkers, whose structural determination immediately suggested that the FH2 dimer “stair-steps” to mediate the sequential incorporation of actin monomers at the barbed-end during processive filament elongation (Xu et al., 2004). Dimerization of the FH2 domain is also necessary for nucleation (Fig. 5C), because each FH2 domain binds an actin monomer such that the ring-like dimer helps stabilize a short-pitch actin dimer (Li and Higgs, 2003; Pruyne et al., 2002). The structure of a formin-actin complex (Otomo et al., 2005) provided further insights into the mechanisms of formin-mediated nucleation and elongation. In the structure, the FH2-actin dimer (built by crystal symmetry) stabilizes a shor-tpitch actin dimer, which could function as a polymerization nucleus. In addition, this FH2 dimer is in contact with a third actin subunit (labeled actin 3 in Fig. 5C), whose position at the barbed end of the filament is supposedly stabilized by a conformational change leading the uppermost subunit of the FH2 dimer (green subunit in Fig. 5C) to move downwards to bind this newly added actin.

Actin filament crosslinking proteins

Spectrin-family proteins, including α-actinin, spectrin, and dystrophin, are the most important group of proteins involved in crosslinking actin filaments, both among them and with cellular organelles and membranes (Broderick and Winder, 2005). These proteins are predominantly localized to the submembrane cytoskeleton, such as the leading edge and focal contacts of migrating cells (Blanchard et al., 1989; Knight et al., 2000; Otto, 1994; Pascual et al., 1997). Actin crosslinking proteins frequently function as molecular scaffolds, connecting actin filament networks to extracellular matrix proteins, such as ankyrin, laminin and dystroglycan (Figs. 2 and and6B)6B) (Bennett and Baines, 2001; Campbell and Kahl, 1989; Kennedy et al., 1991; Rando, 2001).

Fig. 6
Spectrin family of actin crosslinking proteins. (A) Domain organization and structures of a prototypical spectrin-family member. Spectrin-family proteins consist of three regions: an N-terminal actin-binding domain (ABD) formed by two calponin homology ...

Although diverse in size and function, spectrin-family proteins are characterized by the presence of a series of conserved structural modules, including the calponin homology (CH) domain, varying numbers of spectrin repeats, EF-hands, or other regulatory regions (Fig. 6A) (Broderick and Winder, 2005). The CH domain frequently occurs in pairs (CH1 and CH2), which together form the actin-binding domain (ABD) of spectrin-family proteins (Gimona and Winder, 1998).

The crystal structures of several ABDs have been determined, including those of human, A. thaliana and S. pombe fimbrin (Goldsmith et al., 1997; Klein et al., 2004), utrophin (Keep et al., 1999), dystrophin (Norwood et al., 2000), human and mouse plectin (Garcia-Alvarez et al., 2003; Sevcik et al., 2004) and α-actinin 1, 3, 4 (Borrego-Diaz et al., 2006; Franzot et al., 2005; Lee et al., 2008). Generally, these structures display a similar so-called “compact” or “closed” conformation, characterized by extensive interactions between the CH domains. In two of the structures, those of utrophin (Keep et al., 1999) and dystrophin, the compact conformation results from domain swapping between two different molecules in the asymmetric unit (Liu and Eisenberg, 2002). By contrast, the conformation of actin-bound ABDs is debated (Galkin et al., 2002; 2003; Lehman et al., 2004). EM studies of ABD-decorated actin filaments have suggested two different models of binding: compact (Hanein et al., 1998; McGough et al., 1994; Sutherland-Smith et al., 2003) and extended (Galkin et al., 2002; Moores et al., 2000). The compact model holds that the ABD binds actin with only minor changes relative to the crystal structures, whereas the extended model maintains that the two CHs become separated upon binding. It remains to be demonstrated which of these two models is correct.

The spectrin repeats, usually occupy the central region of spectrin-family proteins, and as a result are collectively referred to as the ‘rod’ domain. The number of spectrin repeats (each repeat consisting of ~110-aa) varies significantly among these proteins, ranging from 4 in α-actinin to ~24 in dystrophin (Broderick and Winder, 2005; Pascual et al., 1997). This provides a mechanism for the regulation of the spacing between actin networks and membrane compartments, which can range from 14 to 130 nm (Fig. 6A) (Amann et al., 1999; Broderick and Winder, 2005; Rybakova et al., 2002; Winder et al., 1995). The spectrin- repeat region also mediates anti-parallel dimerization of various spectrin-family proteins. The structures of various spectrin repeat fragments have been determined, including the entire rod domain of α-actinin (Fig. 6A) (Ylanne et al., 2001a; 2001b).

The C-termini of spectrin-family proteins can vary significantly, and distinct types of domains can occur, including the EF-hand and pleckstrin homology (PH) domains, whose specific nature correlates with the individual functions of each protein (Figs. 6A and 6B). For instance, α-actinin, which functions primarily as an actin filament crosslinking protein, contains two C-terminal EF-hand motifs that are thought to come in close contact with the CH domains within the antiparallel dimer, and contribute to modulating the binding of α-actinin to F-actin in a Ca2+- dependent manner (Blanchard et al., 1989; Lundberg et al., 1995; Noegel et al., 1987). On the other hand, β-spectrin, which provides a linkage between actin networks and membranes, presents a c-terminal ph domain, which associates with negatively charged phospholipids at the membrane (Ferguson et al., 1995).

BAR proteins linking membrane and actincytoskeleton dynamics

A tight spatial and temporal coordination of actin polymerization and plasma membrane remodeling is a characteristic feature of many cellular processes, including endocytosis, exocytosis, cell motility and intracellular trafficking (Engqvist-Goldstein and Drubin, 2003; Kaksonen et al., 2006; Scita et al., 2008). In these processes, Bin/Amphiphysin/Rvsp (BAR) domain-containing proteins are emerging as key regulators, linking signaling pathways to actin cytoskeleton and membrane dynamics (Fig. 7) (Dawson et al., 2006; Frost et al., 2007; Itoh and De Camilli, 2006; Scita et al., 2008).

BAR proteins are generally unrelated, but are united by a number of features, including the presence of the membrane-binding BAR domain, their modular organization and the fact that they frequently link to the actin cytoskeleton under the control of Rho-family GTPases (Dawson et al., 2006; Frost et al., 2008; Henne et al., 2007; Itoh and De Camilli, 2006; Scita et al., 2008). Because of the lack of overall sequence similarity, the BAR family has expanded with the determination of crystal structures of the dimerization/membrane-binding domains of proteins originally thought to be unrelated (Habermann, 2004; Millard et al., 2005; Peter et al., 2004; Shimada et al., 2007). The BAR fold consists of an elongated anti-parallel dimer of two helical bundles, with a curved, positively charged membrane-binding interface (Fig. 7A) (Govind et al., 2001; Gallop et al., 2006; Henne et al., 2007; Lee et al., 2007; Masuda et al., 2006; Millard et al., 2005; Peter et al., 2004; Pylypenko et al., 2007; Shimada et al., 2007; Tarricone et al., 2001; Weissenhorn, 2005; Zhu et al., 2007). However, there is significant variation within this fold, and three subfamilies are identified based on the overall shape and curvature of the BAR domain (Frost et al., 2007; Henne et al., 2007; Scita et al., 2008): the classical crescent- shaped N-BAR (found in arfaptin, amphiphysin, and endophilin), the more elongated and less curved F-BAR (found in FBP17, CIP4, FCHo2, and Toca1), and the inverted curvature I-BAR (found in IRSp53, MIM, and ABBA) (Fig. 7A). Some BAR domains present helical appendages (N-BAR) that are thought to penetrate the bound membrane (Gallop et al., 2006; Saarikangas et al., 2009). There is a direct correlation between the shape and size of each BAR domain and the shape and curvature of the membranes associated with them (Fig. 7A) (Frost et al., 2008; Mattila et al., 2007; Peter et al., 2004; Saarikangas et al., 2009; Shimada et al., 2007).

The question of membrane curvature sensing versus generation has received significant attention. While the original view was that BAR proteins primarily ‘senseor’ or ‘stabilize’ curvature (Peter et al., 2004), recent electron microscopy evidence suggests that at least some BAR domains can bind flat membranes and induce curvature via cooperative BAR-BAR interactions that are more favorable on membranes (Frost et al., 2008). This latter study also showed that membrane tubules form when F-BARs polymerize into helical coats that are held together by lateral and tip-to-tip BAR-BAR interactions, as well as interactions of the BAR domain with the membrane itself. Similar interaction may occur with the I-BAR domain, but what distinguished the I-BAR domain is that it induces membrane curvature in the opposite direction to that of the N-BAR and F-vature in the opposite direction to that of the N-BAR and F-BAR domains, and bind to the interior of membrane compartments (Fig. 2) as supposed to the exterior for the N-BAR and F-BAR domains (Mattila et al., 2007; Saarikangas et al., 2009; Suetsugu et al., 2006). The formation of endocytic vesicles is generally driven by BAR proteins that sense membrane curvature and/or actively bend the membrane. Proteins such as Arfarptin and Tuba, containing crescent-shaped membrane-binding BAR domains, sense the curvature of clathrin-coated vesicle, which are ~200 Å in diameter (Salazar et al., 2003; Tarricone et al., 2001). Meanwhile, the N-BAR domains of amphiphysin and endophilin are involved in the formation of the narrowly curved necks of nascent vesicles. These endocytic vesicles are ultimately pinched-off from the membrane by dynamin, a GTPase that catalyzes vesicle scission from the plasma membrane (Fig. 7B) (Gallop et al., 2006; Shafer and Voss, 2004). F-BAR proteins, such as FBP17, syndapin and Toca1, are also implicated in clathrin-mediated endocytosis in cooperation with dynamin (Ho et al., 2004; Kamioka et al., 2004; Kessels and Qualmann, 2004).

The coordination between BAR domain proteins and the actin cytoskeleton is just beginning to be unraveled, but it appears to affect most cellular processes of the actin cytoskeleton. Despite the extraordinary plasticity of the BAR domain, what sets BAR proteins apart is their functional diversity, resulting from the presence of additional modules N- and C-terminal to the BAR domain (Fig. 7B). Thus, a number of BAR proteins present PX or PH domains, which provide added affinity for phospholipid membranes and/or become involved in protein-protein interactions (Pylypenko et al., 2007; Zhu et al., 2007). Many BAR proteins also present modules that link to the actin cytoskeleton, such as WW-binding motifs, SH3 and WH2 domains (Scita et al., 2008; Shimada et al., 2007). Also common is the CRIB domain or other motifs that mediate the binding of Rho GTPases. Most BAR proteins appear to exist in an auto-inhibited conformation, a wide-spread feature among cytoskeletal proteins (Fig. 3C). Thus, a conformational change in IRSp53 resulting from the binding of GTPases to its CRIB motif is thought to free the SH3 domain to recruit ABPs such as WASP/Scar that in turn engage in actin cytoskeleton remodeling (Figs. 3C and and7B)7B) (Govind et al., 2001; Miki et al., 2000; Nakagawa et al., 2003; Takenawa and Suetsugu, 2007). During filopodia and lamellipodia formation, I-BAR proteins are recruited to the membrane and subsequently could direct the barbed ends of the actin filaments toward the plasma membrane by directly interacting with ABPs in response to extracellular stimuli (Ahmed et al., 2009).


The dynamic remodeling of the actin cytoskeleton is an essential component of many cellular processes, including cell locomotion, cytokinesis and membrane trafficking. Additionally, many pathogens hijack the host cell actin cytoskeleton during infection. These processes involve rapid bursts of actin polymerization/ depolymerization with remarkable spatiotemporal precision. Actin and a myriad of actin-binding proteins become involved in the regulation of these processes. A wide range of diseases, including cancer, neurological and musculoskeletal disorders result from malfunctioning of cytoskeletal proteins. While the study of actin cytoskeleton dynamics has recently intensified, what is critically lacking is a comprehensive structure- function understanding of the interplay between the many membrane binding, scaffolding and signaling proteins that conform the cytoskeleton. These questions are likely to dominate the research within the actin cytoskeleton field during the following few years.


This work was supported in part by National Research Foundation (NRF) grants R13-2003-009 through Research Center for Resistant Cells and 2009-0064846 to Sung Haeng Lee, and by NIH grant GM073791 to Roberto Dominguez.


  • Ahmed MA, Bamm VV, Shi L, Steiner-Mosonyi M, Dawson JF, Brown L, Harauz G, Ladizhansky V. Induced secondary structure and polymorphism in an intrinsically disordered structural linker of the CNS: solid-state NMR and FTIR spectroscopy of myelin basic protein bound to actin. Biophys J. 2009;96:180–191. [PubMed]
  • Ahuja R, Pinyol R, Reichenbach N, Custer L, Klingensmith J, Kessels MM, Qualmann B. Cordonbleu is an actin nucleation factor and controls neuronal morphology. Cell. 2007;131:337–350. [PMC free article] [PubMed]
  • Amann KJ, Guo AW, Ervasti JM. Utrophin lacks the rod domain actin binding activity of dystrophin. J Biol Chem. 1999;274:35375–35380. [PubMed]
  • Arber S, Barbayannis FA, Hanser H, Schneider C, Stanyon CA, Bernard O, Caroni P. Regulation of actin dynamics through phosphorylation of cofilin by LIM-kinase. Nature. 1998;393:805–809. [PubMed]
  • Bachmann C, Fischer L, Walter U, Reinhard M. The EVH2 domain of the vasodilator-stimulated phosphoprotein mediates tetramerization, F-actin binding, and actin bundle formation. J Biol Chem. 1999;274:23549–23557. [PubMed]
  • Bailly M, Condeelis J. Cell motility: insights from the backstage. Nat Cell Biol. 2002;4:E292–294. [PubMed]
  • Bamburg JR. Proteins of the ADF/cofilin family: essential regulators of actin dynamics. Annu Rev Cell Dev Biol. 1999;15:185–230. [PubMed]
  • Belmont LD, Orlova A, Drubin DG, Egelman EH. A change in actin conformation associated with filament instability after Pi release. Proc Natl Acad Sci USA. 1999;96:29–34. [PubMed]
  • Bennett V, Baines AJ. Spectrin and ankyrin-based pathways: metazoan inventions for integrating cells into tissues. Physiol Rev. 2001;81:1353–1392. [PubMed]
  • Blanchard A, Ohanian V, Critchley D. The structure and function of alpha-actinin. J Muscle Res Cell Motil. 1989;10:280–289. [PubMed]
  • Borrego-Diaz E, Kerff F, Lee SH, Ferron F, Li Y, Dominguez R. Crystal structure of the actin-binding domain of alpha-actinin 1: evaluating two competing actin-binding models. J Struct Biol. 2006;155:230–238. [PubMed]
  • Bowman GD, Nodelman IM, Hong Y, Chua NH, Lindberg U, Schutt CE. A comparative structural analysis of the ADF/cofilin family. Proteins. 2000;41:374–384. [PubMed]
  • Breitsprecher D, Kiesewetter AK, Linkner J, Urbanke C, Resch GP, Small JV, Faix J. Clustering of VASP actively drives processive, WH2 domain-mediated actin filament elongation. EMBO J. 2008;27:2943–2954. [PubMed]
  • Brindle NP, Holt MR, Davies JE, Price CJ, Critchley DR. The focal-adhesion vasodilator-stimulated phosphoprotein (VASP) binds to the proline-rich domain in vinculin. Biochem J. 1996;318:753–757. [PubMed]
  • Broderick MJ, Winder SJ. Spectrin, alpha-actinin, and dystrophin. Adv Protein Chem. 2005;70:203–246. [PubMed]
  • Burbelo PD, Drechsel D, Hall A. A conserved binding motif defines numerous candidate target proteins for both Cdc42 and Rac GTPases. J Biol Chem. 1995;270:29071–29074. [PubMed]
  • Campbell KP, Kahl SD. Association of dystrophin and an integral membrane glycoprotein. Nature. 1989;338:259–262. [PubMed]
  • Caron E, Hall A. Identification of two distinct mechanisms of phagocytosis controlled by different Rho GTPases. Science. 1998;282:1717–1721. [PubMed]
  • Castrillon DH, Wasserman SA. Diaphanous is required for cytokinesis in Drosophila and shares domains of similarity with the products of the limb deformity gene. Development. 1994;120:3367–3377. [PubMed]
  • Chereau D, Dominguez R. Understanding the role of the G-actin-binding domain of Ena/VASP in actin assembly. J Struct Biol. 2006;155:195–201. [PubMed]
  • Chereau D, Kerff F, Graceffa P, Grabarek Z, Langsetmo K, Dominguez R. Actin-bound structures of Wiskott-Aldrich syndrome protein (WASP)-homology domain 2 and the implications for filament assembly. Proc Natl Acad Sci USA. 2005;102:16644–16649. [PubMed]
  • Chereau D, Boczkowska M, Skwarek-Maruszewska A, Fujiwara I, Hayes DB, Rebowski G, Lappalainen P, Pollard TD, Dominguez R. Leiomodin is an actin filament nucleator in muscle cells. Science. 2008;320:239–243. [PMC free article] [PubMed]
  • Condeelis J, Singer RH, Segall JE. The great escape: when cancer cells hijack the genes for chemotaxis and motility. Annu Rev Cell Dev Biol. 2005;21:695–718. [PubMed]
  • Dawson JC, Legg JA, Machesky LM. Bar domain proteins: a role in tubulation, scission and actin assembly in clathrin-mediated endocytosis. Trends Cell Biol. 2006;16:493–498. [PubMed]
  • Doctor RB, Fouassier L. Emerging roles of the actin cytoskeleton in cholangiocyte function and disease. Semin Liver Dis. 2002;22:263–276. [PubMed]
  • Domanski M, Hertzog M, Coutant J, Gutsche-Perelroizen I, Bontems F, Carlier MF, Guittet E, van Heijenoort C. Coupling of folding and binding of thymosin beta4 upon interaction with monomeric actin monitored by nuclear magnetic resonance. J Biol Chem. 2004;279:23637–23645. [PubMed]
  • Dominguez R. Actin-binding proteins--a unifying hypothesis. Trends Biochem Sci. 2004;29:572–578. [PubMed]
  • Dominguez R. The beta-thymosin/WH2 fold: multifunctionality and structure. Ann N Y Acad Sci. 2007;1112:86–94. [PubMed]
  • Dominguez R. Actin filament nucleation and elongation factors--structure-function relationships. Crit Rev Biochem Mol Biol. 2009;44:351–366. [PMC free article] [PubMed]
  • Edwards DC, Sanders LC, Bokoch GM, Gill GN. Activation of LIM-kinase by Pak1 couples Rac/Cdc42 GTPase signalling to actin cytoskeletal dynamics. Nat Cell Biol. 1999;1:253–259. [PubMed]
  • Egelman EH. Actin filament structure. The ghost of ribbons past. Curr Biol. 1994;4:79–81. [PubMed]
  • Eisenmann KM, Harris ES, Kitchen SM, Holman HA, Higgs HN, Alberts AS. Dia-interacting protein modulates formin-mediated actin assembly at the cell cortex. Curr Biol. 2007;17:579–591. [PubMed]
  • Engqvist-Goldstein AE, Drubin DG. Actin assembly and endocytosis: from yeast to mammals. Annu Rev Cell Dev Biol. 2003;19:287–332. [PubMed]
  • Etienne-Manneville S, Hall A. Rho GTPases in cell biology. Nature. 2002;420:629–635. [PubMed]
  • Fedorov AA, Lappalainen P, Fedorov EV, Drubin DG, Almo SC. Structure determination of yeast cofilin. Nat Struct Biol. 1997;4:366–369. [PubMed]
  • Ferguson KM, Lemmon MA, Sigler PB, Schlessinger J. Scratching the surface with the PH domain. Nat Struct Biol. 1995;2:715–718. [PubMed]
  • Ferron F, Rebowski G, Lee SH, Dominguez R. Structural basis for the recruitment of profilin-actin complexes during filament elongation by Ena/VASP. EMBO J. 2007;26:4597–4606. [PubMed]
  • Foster R, Hu KQ, Lu Y, Nolan KM, Thissen J, Settleman J. Identification of a novel human Rho protein with unusual properties: GTPase deficiency and in vivo farnesylation. Mol Cell Biol. 1996;16:2689–2699. [PMC free article] [PubMed]
  • Franzot G, Sjoblom B, Gautel M, Djinovic Carugo K. The crystal structure of the actin binding domain from alpha-actinin in its closed conformation: structural insight into phospholipid regulation of alpha-actinin. J Mol Biol. 2005;348:151–165. [PubMed]
  • Frost A, De Camilli P, Unger VM. F-BAR proteins join the BAR family fold. Structure. 2007;15:751–753. [PubMed]
  • Frost A, Perera R, Roux A, Spasov K, Destaing O, Egelman EH, De Camilli P, Unger VM. Structural basis of membrane invagination by F-BAR domains. Cell. 2008;132:807–817. [PMC free article] [PubMed]
  • Galkin VE, Orlova A, VanLoock MS, Rybakova IN, Ervasti JM, Egelman EH. The utrophin actin-binding domain binds F-actin in two different modes: implications for the spectrin superfamily of proteins. J Cell Biol. 2002;157:243–251. [PMC free article] [PubMed]
  • Galkin VE, Orlova A, VanLoock MS, Egelman EH. Do the utrophin tandem calponin homology domains bind F-actin in a compact or extended conformation? J Mol Biol. 2003;331:967–972. [PubMed]
  • Gallop JL, Jao CC, Kent HM, Butler PJ, Evans PR, Langen R, McMahon HT. Mechanism of endophilin N-BAR domain-mediated membrane curvature. EMBO J. 2006;25:2898–2910. [PubMed]
  • Garcia-Alvarez B, Bobkov A, Sonnenberg A, de Pereda JM. Structural and functional analysis of the actin binding domain of plectin suggests alternative mechanisms for binding to F-actin and integrin beta4. Structure. 2003;11:615–625. [PubMed]
  • Gimona M, Djinovic-Carugo K, Kranewitter WJ, Winder SJ. Functional plasticity of CH domains. FEBS Lett. 2002;513:98–106. [PubMed]
  • Gimona M, Winder SJ. Single calponin homology domains are not actin-binding domains. Curr Biol. 1998;8:R674–675. [PubMed]
  • Goldsmith SC, Pokala N, Shen W, Fedorov AA, Matsudaira P, Almo SC. The structure of an actin-crosslinking domain from human fimbrin. Nat Struct Biol. 1997;4:708–712. [PubMed]
  • Goode BL, Eck MJ. Mechanism and function of formins in the control of actin assembly. Annu Rev Biochem. 2007;76:593–627. [PubMed]
  • Govind S, Kozma R, Monfries C, Lim L, Ahmed S. Cdc42Hs facilitates cytoskeletal reorganization and neurite outgrowth by localizing the 58-kD insulin receptor substrate to filamentous actin. J Cell Biol. 2001;152:579–594. [PMC free article] [PubMed]
  • Graceffa P, Dominguez R. Crystal structure of monomeric actin in the ATP state. Structural basis of nucleotide-dependent actin dynamics. J Biol Chem. 2003;278:34172–34180. [PubMed]
  • Grintsevich EE, Benchaar SA, Warshaviak D, Boontheung P, Halgand F, Whitelegge JP, Faull KF, Loo RR, Sept D, Loo JA, et al. Mapping the cofilin binding site on yeast G-actin by chemical cross-linking. J Mol Biol. 2008;377:395–409. [PMC free article] [PubMed]
  • Guan JQ, Vorobiev S, Almo SC, Chance MR. Mapping the G-actin binding surface of cofilin using synchrotron protein footprinting. Biochemistry. 2002;41:5765–5775. [PubMed]
  • Habermann B. The BAR-domain family of proteins: a case of bending and binding? EMBO Rep. 2004;5:250–255. [PubMed]
  • Hall A. Small GTP-binding proteins and the regulation of the actin cytoskeleton. Annu Rev Cell Biol. 1994;10:31–54. [PubMed]
  • Hanein D, Volkmann N, Goldsmith S, Michon AM, Lehman W, Craig R, DeRosier D, Almo S, Matsudaira P. An atomic model of fimbrin binding to F-actin and its implications for filament crosslinking and regulation. Nat Struct Biol. 1998;5:787–792. [PubMed]
  • Helfer E, Nevalainen EM, Naumanen P, Romero S, Didry D, Pantaloni D, Lappalainen P, Carlier MF. Mammalian twinfilin sequesters ADP-G-actin and caps filament barbed ends: implications in motility. EMBO J. 2006;25:1184–1195. [PubMed]
  • Henne WM, Kent HM, Ford MG, Hegde BG, Daumke O, Butler PJ, Mittal R, Langen R, Evans PR, McMahon HT. Structure and analysis of FCHo2 F-BAR domain: a dimerizing and membrane recruitment module that effects membrane curvature. Structure. 2007;15:839–852. [PubMed]
  • Hertzog M, van Heijenoort C, Didry D, Gaudier M, Coutant J, Gigant B, Didelot G, Preat T, Knossow M, Guittet E, et al. The beta-thymosin/WH2 domain; structural basis for the switch from inhibition to promotion of actin assembly. Cell. 2004;117:611–623. [PubMed]
  • Ho HY, Rohatgi R, Lebensohn AM, Le M, Li J, Gygi SP, Kirschner MW. Toca-1 mediates Cdc42-dependent actin nucleation by activating the N-WASP-WIP complex. Cell. 2004;118:203–216. [PubMed]
  • Holmes KC, Popp D, Gebhard W, Kabsch W. Atomic model of the actin filament. Nature. 1990;347:44–49. [PubMed]
  • Huang KY, Lai MW, Lee WI, Huang YC. Fatal cytomegalovirus gastrointestinal disease in an infant with Wiskott-Aldrich syndrome. J Formos Med Assoc. 2008;107:64–67. [PubMed]
  • Hussey PJ, Ketelaar T, Deeks MJ. Control of the actin cytoskeleton in plant cell growth. Annu Rev Plant Biol. 2006;57:109–125. [PubMed]
  • Itoh T, De Camilli P. BAR, F-BAR (EFC) and ENTH/ ANTH domains in the regulation of membrane-cytosol interfaces and membrane curvature. Biochim Biophys Acta. 2006;1761:897–912. [PubMed]
  • Itoh G, Yumura S. A novel mitosis-specific dynamic actin structure in Dictyostelium cells. J Cell Sci. 2007;120:4302–4309. [PubMed]
  • Janmey PA, Stossel TP, Lind SE. Sequential binding of actin monomers to plasma gelsolin and its inhibition by vitamin D-binding protein. Biochem Biophys Res Commun. 1986;136:72–79. [PubMed]
  • Kaksonen M, Toret CP, Drubin DG. Harnessing actin dynamics for clathrin-mediated endocytosis. Nat Rev Mol Cell Biol. 2006;7:404–414. [PubMed]
  • Kamioka Y, Fukuhara S, Sawa H, Nagashima K, Masuda M, Matsuda M, Mochizuki N. A novel dynamin-associating molecule, formin-binding protein 17, induces tubular membrane invaginations and participates in endocytosis. J Biol Chem. 2004;279:40091–40099. [PubMed]
  • Keep NH, Winder SJ, Moores CA, Walke S, Norwood FL, Kendrick-Jones J. Crystal structure of the actin-binding region of utrophin reveals a head-to-tail dimer. Structure. 1999;7:1539–1546. [PubMed]
  • Kennedy SP, Warren SL, Forget BG, Morrow JS. Ankyrin binds to the 15th repetitive unit of erythroid and nonerythroid beta-spectrin. J Cell Biol. 1991;115:267–277. [PMC free article] [PubMed]
  • Kessels MM, Qualmann B. The syndapin protein family: linking membrane trafficking with the cytoskeleton. J Cell Sci. 2004;117:3077–3086. [PubMed]
  • Klein MG, Shi W, Ramagopal U, Tseng Y, Wirtz D, Kovar DR, Staiger CJ, Almo SC. Structure of the actin crosslinking core of fimbrin. Structure. 2004;12:999–1013. [PubMed]
  • Knight B, Laukaitis C, Akhtar N, Hotchin NA, Edlund M, Horwitz AR. Visualizing muscle cell migration in situ. Curr Biol. 2000;10:576–585. [PubMed]
  • Kovar DR, Harris ES, Mahaffy R, Higgs HN, Pollard TD. Control of the assembly of ATP- and ADP-actin by formins and profilin. Cell. 2006;124:423–435. [PubMed]
  • Kozma R, Ahmed S, Best A, Lim L. The Ras-related protein Cdc42Hs and bradykinin promote formation of peripheral actin microspikes and filopodia in Swiss 3T3 fibroblasts. Mol Cell Biol. 1995;15:1942–1952. [PMC free article] [PubMed]
  • Krause M, Dent EW, Bear JE, Loureiro JJ, Gertler FB. Ena/VASP proteins: regulators of the actin cytoskeleton and cell migration. Annu Rev Cell Dev Biol. 2003;19:541–564. [PubMed]
  • Krause M, Leslie JD, Stewart M, Lafuente EM, Valderrama F, Jagannathan R, Strasser GA, Rubinson DA, Liu H, Way M, et al. Lamellipodin, an Ena/VASP ligand, is implicated in the regulation of lamellipodial dynamics. Dev Cell. 2004;7:571–583. [PubMed]
  • Krugmann S, Jordens I, Gevaert K, Driessens M, Vandekerckhove J, Hall A. Cdc42 induces filopodia by promoting the formation of an IRSp53:Mena complex. Curr Biol. 2001;11:1645–1655. [PubMed]
  • Kuhnel K, Jarchau T, Wolf E, Schlichting I, Walter U, Wittinghofer A, Strelkov SV. The VASP tetramerization domain is a right-handed coiled coil based on a 15-residue repeat. Proc Natl Acad Sci USA. 2004;101:17027–17032. [PubMed]
  • Lappalainen P, Fedorov EV, Fedorov AA, Almo SC, Drubin DG. Essential functions and actin-binding surfaces of yeast cofilin revealed by systematic mutagenesis. EMBO J. 1997;16:5520–5530. [PubMed]
  • Lappalainen P, Kessels MM, Cope MJ, Drubin DG. The ADF homology (ADF-H) domain: a highly exploited actin-binding module. Mol Biol Cell. 1998;9:1951–1959. [PMC free article] [PubMed]
  • Lee SH, Kerff F, Chereau D, Ferron F, Klug A, Dominguez R. Structural basis for the actin-binding function of missing-in-metastasis. Structure. 2007;15:145–155. [PMC free article] [PubMed]
  • Lee SH, Weins A, Hayes DB, Pollak MR, Dominguez R. Crystal structure of the actin-binding domain of alpha-actinin- 4 Lys255Glu mutant implicated in focal segmental glomerulosclerosis. J Mol Biol. 2008;376:317–324. [PMC free article] [PubMed]
  • Lehman W, Craig R, Kendrick-Jones J, Sutherland-Smith AJ. An open or closed case for the conformation of calponin homology domains on F-actin? J Muscle Res Cell Motil. 2004;25:351–358. [PubMed]
  • Li F, Higgs HN. The mouse Formin mDia1 is a potent actin nucleation factor regulated by autoinhibition. Curr Biol. 2003;13:1335–1340. [PubMed]
  • Liu Y, Eisenberg D. 3D domain swapping: as domains continue to swap. Protein Sci. 2002;11:1285–1299. [PubMed]
  • Liverman AD, Cheng HC, Trosky JE, Leung DW, Yarbrough ML, Burdette DL, Rosen MK, Orth K. Arp2/3-independent assembly of actin by Vibrio type III effector VopL. Proc Natl Acad Sci USA. 2007;104:17117–17122. [PubMed]
  • Lu Y, Settleman J. The role of rho family GTPases in development: lessons from Drosophila melanogaster. Mol Cell Biol Res Commun. 1999;1:87–94. [PubMed]
  • Lundberg S, Bjork J, Lofvenberg L, Backman L. Cloning, expression and characterization of two putative calcium-binding sites in human non-erythroid alpha-spectrin. Eur J Biochem. 1995;230:658–665. [PubMed]
  • Luo L, Jan LY, Jan YN. Rho family GTP-binding proteins in growth cone signalling. Curr Opin Neurobiol. 1997;7:81–86. [PubMed]
  • Mabuchi I, Hamaguchi Y, Fujimoto H, Morii N, Mishima M, Narumiya S. A rho-like protein is involved in the organisation of the contractile ring in dividing sand dollar eggs. Zygote. 1993;1:325–331. [PubMed]
  • Machesky LM, Gould KL. The Arp2/3 complex: a multifunctional actin organizer. Curr Opin Cell Biol. 1999;11:117–121. [PubMed]
  • Machesky LM, Insall RH. Scar1 and the related Wiskott-Aldrich syndrome protein, WASP, regulate the actin cytoskeleton through the Arp2/3 complex. Curr Biol. 1998;8:1347–1356. [PubMed]
  • Machesky LM, Atkinson SJ, Ampe C, Vandekerckhove J, Pollard TD. Purification of a cortical complex containing two unconventional actins from Acanthamoeba by affinity chromatography on profilin-agarose. J Cell Biol. 1994;127:107–115. [PMC free article] [PubMed]
  • Machesky LM, Mullins RD, Higgs HN, Kaiser DA, Blanchoin L, May RC, Hall ME, Pollard TD. Scar, a WASp-related protein, activates nucleation of actin filaments by the Arp2/3 complex. Proc Natl Acad Sci USA. 1999;96:3739–3744. [PubMed]
  • Machner MP, Urbanke C, Barzik M, Otten S, Sechi AS, Wehland J, Heinz DW. ActA from Listeria mono-cytogenes can interact with up to four Ena/VASP homology 1 domains simultaneously. J Biol Chem. 2001;276:40096–40103. [PubMed]
  • Maekawa M, Ishizaki T, Boku S, Watanabe N, Fujita A, Iwamatsu A, Obinata T, Ohashi K, Mizuno K, Narumiya S. Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase. Science. 1999;285:895–898. [PubMed]
  • Masuda M, Takeda S, Sone M, Ohki T, Mori H, Kamioka Y, Mochizuki N. Endophilin BAR domain drives membrane curvature by two newly identified structure-based mechanisms. EMBO J. 2006;25:2889–2897. [PubMed]
  • Mattila PK, Lappalainen P. Filopodia: molecular architecture and cellular functions. Nat Rev Mol Cell Biol. 2008;9:446–454. [PubMed]
  • Mattila PK, Pykalainen A, Saarikangas J, Paavilainen VO, Vihinen H, Jokitalo E, Lappalainen P. Missing-in-metastasis and IRSp53 deform PI(4,5)P2-rich membranes by an inverse BAR domain-like mechanism. J Cell Biol. 2007;176:953–964. [PMC free article] [PubMed]
  • McGough A, Way M, DeRosier D. Determination of the alpha-actinin-binding site on actin filaments by cryoelectron microscopy and image analysis. J Cell Biol. 1994;126:433–443. [PMC free article] [PubMed]
  • McGough AM, Staiger CJ, Min JK, Simonetti KD. The gelsolin family of actin regulatory proteins: modular structures, versatile functions. FEBS Lett. 2003;552:75–81. [PubMed]
  • McLaughlin PJ, Gooch JT, Mannherz HG, Weeds AG. Structure of gelsolin segment 1-actin complex and the mechanism of filament severing. Nature. 1993;364:685–692. [PubMed]
  • McMahon HT, Gallop JL. Membrane curvature and mechanisms of dynamic cell membrane remodelling. Nature. 2005;438:590–596. [PubMed]
  • Miki H, Yamaguchi H, Suetsugu S, Takenawa T. IRSp53 is an essential intermediate between Rac and WAVE in the regulation of membrane ruffling. Nature. 2000;408:732–735. [PubMed]
  • Millard TH, Bompard G, Heung MY, Dafforn TR, Scott DJ, Machesky LM, Futterer K. Structural basis of filopodia formation induced by the IRSp53/MIM homology domain of human IRSp53. EMBO J. 2005;24:240–250. [PubMed]
  • Moores CA, Keep NH, Kendrick-Jones J. Structure of the utrophin actin-binding domain bound to F-actin reveals binding by an induced fit mechanism. J Mol Biol. 2000;297:465–480. [PubMed]
  • Mullins RD, Heuser JA, Pollard TD. The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc Natl Acad Sci USA. 1998;95:6181–6186. [PubMed]
  • Myers KA, He Y, Hasaka TP, Baas PW. Microtubule transport in the axon: re-thinking a potential role for the actin cytoskeleton. Neuroscientist. 2006;12:107–118. [PubMed]
  • Nakagawa H, Miki H, Nozumi M, Takenawa T, Miyamoto S, Wehland J, Small JV. IRSp53 is colocalised with WAVE2 at the tips of protruding lamellipodia and filopodia independently of Mena. J Cell Sci. 2003;116:2577–2583. [PubMed]
  • Nobes CD, Hall A. Rho GTPases control polarity, protrusion, and adhesion during cell movement. J Cell Biol. 1999;144:1235–1244. [PMC free article] [PubMed]
  • Noegel A, Witke W, Schleicher M. Calcium-sensitive non-muscle alpha-actinin contains EF-hand structures and highly conserved regions. FEBS Lett. 1987;221:391–396. [PubMed]
  • Norwood FL, Sutherland-Smith AJ, Keep NH, Kendrick-Jones J. The structure of the N-terminal actin-binding domain of human dystrophin and how mutations in this domain may cause Duchenne or Becker muscular dystrophy. Structure. 2000;8:481–491. [PubMed]
  • Oda T, Iwasa M, Aihara T, Maeda Y, Narita A. The nature of the globular- to fibrous-actin transition. Nature. 2009;457:441–445. [PubMed]
  • Ojala PJ, Paavilainen VO, Vartiainen MK, Tuma R, Weeds AG, Lappalainen P. The two ADF-H domains of twinfilin play functionally distinct roles in interactions with actin monomers. Mol Biol Cell. 2002;13:3811–3821. [PMC free article] [PubMed]
  • Otomo T, Tomchick DR, Otomo C, Panchal SC, Machius M, Rosen MK. Structural basis of actin filament nucleation and processive capping by a formin homology 2 domain. Nature. 2005;433:488–494. [PubMed]
  • Otterbein LR, Graceffa P, Dominguez R. The crystal structure of uncomplexed actin in the ADP state. Science. 2001;293:708–711. [PubMed]
  • Otterbein LR, Cosio C, Graceffa P, Dominguez R. Crystal structures of the vitamin D-binding protein and its complex with actin: structural basis of the actin-scavenger system. Proc Natl Acad Sci USA. 2002;99:8003–8008. [PubMed]
  • Otto JJ. Actin-bundling proteins. Curr Opin Cell Biol. 1994;6:105–109. [PubMed]
  • Paavilainen VO, Merckel MC, Falck S, Ojala PJ, Pohl E, Wilmanns M, Lappalainen P. Structural conservation between the actin monomer-binding sites of twinfilin and actin-depolymerizing factor (ADF)/cofilin. J Biol Chem. 2002;277:43089–43095. [PubMed]
  • Paavilainen VO, Oksanen E, Goldman A, Lappalainen P. Structure of the actin-depolymerizing factor homology domain in complex with actin. J Cell Biol. 2008;182:51–59. [PMC free article] [PubMed]
  • Pascual J, Pfuhl M, Walther D, Saraste M, Nilges M. Solution structure of the spectrin repeat: a left-handed antiparallel triple-helical coiled-coil. J Mol Biol. 1997;273:740–751. [PubMed]
  • Pasic L, Kotova T, Schafer DA. Ena/VASP proteins capture actin filament barbed ends. J Biol Chem. 2008;283:9814–9819. [PMC free article] [PubMed]
  • Paunola E, Mattila PK, Lappalainen P. WH2 domain: a small, versatile adapter for actin monomers. FEBS Lett. 2002;513:92–97. [PubMed]
  • Peng J, Wallar BJ, Flanders A, Swiatek PJ, Alberts AS. Disruption of the Diaphanous-related formin Drf1 gene encoding mDia1 reveals a role for Drf3 as an effector for Cdc42. Curr Biol. 2003;13:534–545. [PubMed]
  • Peter BJ, Kent HM, Mills IG, Vallis Y, Butler PJ, Evans PR, McMahon HT. BAR domains as sensors of membrane curvature: the amphiphysin BAR structure. Science. 2004;303:495–499. [PubMed]
  • Pollard TD, Borisy GG. Cellular motility driven by assembly and disassembly of actin filaments. Cell. 2003;112:453–465. [PubMed]
  • Pollard TD, Blanchoin L, Mullins RD. Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu Rev Biophys Biomol Struct. 2000;29:545–576. [PubMed]
  • Prehoda KE, Scott JA, Mullins RD, Lim WA. Integration of multiple signals through cooperative regulation of the N-WASP-Arp2/3 complex. Science. 2000;290:801–806. [PubMed]
  • Prokopenko SN, Saint R, Bellen HJ. Untying the Gordian knot of cytokinesis. Role of small G proteins and their regulators. J Cell Biol. 2000;148:843–848. [PMC free article] [PubMed]
  • Pruyne D, Evangelista M, Yang C, Bi E, Zigmond S, Bretscher A, Boone C. Role of formins in actin assembly: nucleation and barbed-end association. Science. 2002;297:612–615. [PubMed]
  • Puppo A, Chun JT, Gragnaniello G, Garante E, Santella L. Alteration of the cortical actin cytoskeleton deregulates Ca2+ signaling, monospermic fertilization, and sperm entry. PLoS ONE. 2008;3:e3588. [PMC free article] [PubMed]
  • Pylypenko O, Lundmark R, Rasmuson E, Carlsson SR, Rak A. The PX-BAR membrane-remodeling unit of sorting nexin 9. EMBO J. 2007;26:4788–4800. [PubMed]
  • Quinlan ME, Heuser JE, Kerkhoff E, Mullins RD. Drosophila Spire is an actin nucleation factor. Nature. 2005;433:382–388. [PubMed]
  • Rafelski SM, Theriot JA. Crawling toward a unified model of cell mobility: spatial and temporal regulation of actin dynamics. Annu Rev Biochem. 2004;73:209–239. [PubMed]
  • Raftopoulou M, Hall A. Cell migration: Rho GTPases lead the way. Dev Biol. 2004;265:23–32. [PubMed]
  • Rando TA. The dystrophin-glycoprotein complex, cellular signaling, and the regulation of cell survival in the muscular dystrophies. Muscle Nerve. 2001;24:1575–1594. [PubMed]
  • Renault L, Bugyi B, Carlier MF. Spire and Cordonbleu: multifunctional regulators of actin dynamics. Trends Cell Biol. 2008;18:494–504. [PubMed]
  • Ridley AJ, Hall A. The small GTP-binding protein rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell. 1992;70:389–399. [PubMed]
  • Ridley AJ, Paterson HF, Johnston CL, Diekmann D, Hall A. The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell. 1992;70:401–410. [PubMed]
  • Robinson RC, Turbedsky K, Kaiser DA, Marchand JB, Higgs HN, Choe S, Pollard TD. Crystal structure of Arp2/3 complex. Science. 2001;294:1679–1684. [PubMed]
  • Rybakova IN, Patel JR, Davies KE, Yurchenco PD, Ervasti JM. Utrophin binds laterally along actin filaments and can couple costameric actin with sarcolemma when over-expressed in dystrophin-deficient muscle. Mol Biol Cell. 2002;13:1512–1521. [PMC free article] [PubMed]
  • Saarikangas J, Zhao H, Pykalainen A, Laurinmaki P, Mattila PK, Kinnunen PK, Butcher SJ, Lappalainen P. Molecular mechanisms of membrane deformation by I-BAR domain proteins. Curr Biol. 2009;19:95–107. [PubMed]
  • Salazar MA, Kwiatkowski AV, Pellegrini L, Cestra G, Butler MH, Rossman KL, Serna DM, Sondek J, Gertler FB, De Camilli P. Tuba, a novel protein containing bin/ amphiphysin/Rvs and Dbl homology domains, links dynamin to regulation of the actin cytoskeleton. J Biol Chem. 2003;278:49031–49043. [PubMed]
  • Schutt CE, Myslik JC, Rozycki MD, Goonesekere NC, Lindberg U. The structure of crystalline profilin-beta-actin. Nature. 1993;365:810–816. [PubMed]
  • Scita G, Confalonieri S, Lappalainen P, Suetsugu S. IRSp53: crossing the road of membrane and actin dynamics in the formation of membrane protrusions. Trends Cell Biol. 2008;18:52–60. [PubMed]
  • Scoville D, Stamm JD, Toledo-Warshaviak D, Altenbach C, Phillips M, Shvetsov A, Rubenstein PA, Hubbell WL, Reisler E. Hydrophobic loop dynamics and actin filament stability. Biochemistry. 2006;45:13576–13584. [PubMed]
  • Sellers JR. Myosins: a diverse superfamily. Biochim Biophys Acta. 2000;1496:3–22. [PubMed]
  • Settleman J. Rho GTPases in development. Prog Mol Subcell Biol. 1999;22:201–229. [PubMed]
  • Sevcik J, Urbanikova L, Kost’an J, Janda L, Wiche G. Actin-binding domain of mouse plectin. Crystal structure and binding to vimentin. Eur J Biochem. 2004;271:1873–1884. [PubMed]
  • Shafer A, Voss J. The use of spin-labeled ligands as biophysical probes to report real-time endocytosis of G protein-coupled receptors in living cells. Sci STKE. 2004;2004:pl9. [PubMed]
  • Shimada A, Niwa H, Tsujita K, Suetsugu S, Nitta K, Hanawa-Suetsugu K, Akasaka R, Nishino Y, Toyama M, Chen L, et al. Curved EFC/F-BAR-domain dimers are joined end to end into a filament for membrane invagination in endocytosis. Cell. 2007;129:761–772. [PubMed]
  • Suetsugu S, Murayama K, Sakamoto A, Hanawa-Suetsugu K, Seto A, Oikawa T, Mishima C, Shirouzu M, Takenawa T, Yokoyama S. The RAC binding domain/IRSp53-MIM homology domain of IRSp53 induces RAC-dependent membrane deformation. J Biol Chem. 2006;281:35347–35358. [PubMed]
  • Sutherland-Smith AJ, Moores CA, Norwood FL, Hatch V, Craig R, Kendrick-Jones J, Lehman W. An atomic model for actin binding by the CH domains and spectrin-repeat modules of utrophin and dystrophin. J Mol Biol. 2003;329:15–33. [PubMed]
  • Takenawa T, Suetsugu S. The WASP-WAVE protein network: connecting the membrane to the cytoskeleton. Nat Rev Mol Cell Biol. 2007;8:37–48. [PubMed]
  • Tam VC, Serruto D, Dziejman M, Brieher W, Mekalanos JJ. A type III secretion system in Vibrio cholerae translocates a formin/spire hybrid-like actin nucleator to promote intestinal colonization. Cell Host Microbe. 2007;1:95–107. [PubMed]
  • Tarricone C, Xiao B, Justin N, Walker PA, Rittinger K, Gamblin SJ, Smerdon SJ. The structural basis of Arfaptin-mediated cross-talk between Rac and Arf signalling pathways. Nature. 2001;411:215–219. [PubMed]
  • Vartiainen MK, Guettler S, Larijani B, Treisman R. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science. 2007;316:1749–1752. [PubMed]
  • Watanabe N, Madaule P, Reid T, Ishizaki T, Watanabe G, Kakizuka A, Saito Y, Nakao K, Jockusch BM, Narumiya S. p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J. 1997;16:3044–3056. [PubMed]
  • Weissenhorn W. Crystal structure of the endophilin-A1 BAR domain. J Mol Biol. 2005;351:653–661. [PubMed]
  • Winder SJ, Gibson TJ, Kendrick-Jones J. Dystrophin and utrophin: the missing links! FEBS Lett. 1995;369:27–33. [PubMed]
  • Xu Y, Moseley JB, Sagot I, Poy F, Pellman D, Goode BL, Eck MJ. Crystal structures of a Formin Homology-2 domain reveal a tethered dimer architecture. Cell. 2004;116:711–723. [PubMed]
  • Yamaguchi H, Condeelis J. Regulation of the actin cytoskeleton in cancer cell migration and invasion. Biochim Biophys Acta. 2007;1773:642–652. [PMC free article] [PubMed]
  • Yamagishi A, Masuda M, Ohki T, Onishi H, Mochizuki N. A novel actin bundling/filopodium-forming domain conserved in insulin receptor tyrosine kinase substrate p53 and missing in metastasis protein. J Biol Chem. 2004;279:14929–14936. [PubMed]
  • Yang N, Higuchi O, Ohashi K, Nagata K, Wada A, Kangawa K, Nishida E, Mizuno K. Cofilin phosphorylation by LIM-kinase 1 and its role in Rac-mediated actin reorganization. Nature. 1998;393:809–812. [PubMed]
  • Yarmola EG, Parikh S, Bubb MR. Formation and implications of a ternary complex of profilin, thymosin beta 4, and actin. J Biol Chem. 2001;276:45555–45563. [PubMed]
  • Yarmola EG, Klimenko ES, Fujita G, Bubb MR. Thymosin beta4: actin regulation and more. Ann N Y Acad Sci. 2007;1112:76–85. [PubMed]
  • Ylanne J, Scheffzek K, Young P, Saraste M. Crystal structure of the alpha-actinin rod reveals an extensive torsional twist. Structure. 2001a;9:597–604. [PubMed]
  • Ylanne J, Scheffzek K, Young P, Saraste M. Crystal structure of the alpha-actinin rod: four spectrin repeats forming a thight dimer. Cell Mol Biol Lett. 2001b;6:234. [PubMed]
  • Zhu G, Chen J, Liu J, Brunzelle JS, Huang B, Wakeham N, Terzyan S, Li X, Rao Z, Li G, et al. Structure of the APPL1 BAR-PH domain and characterization of its interaction with Rab5. EMBO J. 2007;26:3484–3493. [PMC free article] [PubMed]
  • Zigmond SH. Beginning and ending an actin filament: control at the barbed end. Curr Top Dev Biol. 2004a;63:145–188. [PubMed]
  • Zigmond SH. Formin-induced nucleation of actin filaments. Curr Opin Cell Biol. 2004b;16:99–105. [PubMed]