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Mechanisms of actin disassembly and turnover

Bruce L. Goode, Julian Eskin, Shashank Shekhar

Journal of Cell Biology · 2023 · Review · CC BY

doi:10.1083/jcb.202309021

Abstract

Cellular actin networks exhibit a wide range of sizes, shapes, and architectures tailored to their biological roles. Once assembled, these filamentous networks are either maintained in a state of polarized turnover or induced to undergo net disassembly. Further, the rates at which the networks are turned over and/or dismantled can vary greatly, from seconds to minutes to hours or even days. Here, we review the molecular machinery and mechanisms employed in cells to drive the disassembly and turnover of actin networks. In particular, we highlight recent discoveries showing that specific combinations of conserved actin disassembly-promoting proteins (cofilin, GMF, twinfilin, Srv2/CAP, coronin, AIP1, capping protein, and profilin) work in concert to debranch, sever, cap, and depolymerize actin filaments, and to recharge actin monomers for new rounds of assembly.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 27, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Actin is one of the most abundant proteins in eukaryotic cells (>100 µM in vertebrate cells) and readily polymerizes into filaments. Actin filaments are spatially organized into elaborate networks with specific biological functions, ranging from muscle contraction and cell migration to endocytosis, intracellular transport, and cytokinesis. Depending on its cellular function, actin networks adopt highly variable shapes, sizes, and filamentous architectures. This includes branched actin arrays found at the cell’s leading edge and sites of endocytosis, tightly packed parallel bundles in filopodia and stereocilia, and antiparallel arrays in muscle sarcomeres, stress fibers, and cytokinetic actin rings (Fig. 1).

In the eukaryotic cytosol, a pool of actin monomers is maintained at high concentrations (10–200 µM), orders of magnitude above the critical concentration for filament assembly (0.12 µM; ). This is achieved by a combination of cellular factors, including high concentrations of profilin and thymosin-β4 to bind monomers and capping protein (CP) to limit growth at filament barbed ends. By capping the majority of barbed ends in the cell, the pool of actin monomers builds up to high levels and is “funneled” to the small fraction of barbed ends that are uncapped, feeding their rapid growth.

Cytosolic conditions strongly favor actin assembly; therefore, specialized mechanisms are needed to destabilize filaments and drive their disassembly. Indeed, dynamic turnover of actin networks is crucial for their biological functions and gives cells the plasticity necessary to respond to signaling events and rapidly rearrange and/or dismantle actin networks. Turnover refers to the polarized flux of actin subunits through a filamentous actin network, where subunits are preferentially added at one end of the network and lost at the other. This polarity reflects the polarity of individual actin filaments, which are believed to assemble predominantly at the “barbed ends” and depolymerize at the “pointed ends” (Box 1). Filaments assembled from purified actin turn over slowly in vitro (one subunit every 3–4 s). However, pioneering work from Yu-li Wang showed that F-actin networks in living cells turn over orders of magnitude faster, suggesting that cells must have factors that catalyze actin disassembly (Box 2). In vivo, the turnover rates of different actin structures can vary greatly, from seconds for the networks at the leading edge and sites of endocytosis to hours or even days for stereocilia and sarcomeres (;;; Fig. 1). How such a wide range of turnover rates is specified in vivo is only now beginning to be understood and appears to involve differential tuning of the actin disassembly mechanisms described below.

A unifying feature of actin networks is their polarity, which stems from the polarity of their constituent actin filaments. Actin filaments have a barbed end, where most of the growth occurs in vivo, and a pointed end, where most of the disassembly is thought to occur. The barbed and pointed ends of the filament have highly distinct kinetic properties, which now have been explained by important differences in the conformation of actin subunits at the two ends, as revealed by a high-resolution cryoEM structure of F-actin. The changing nucleotide state of actin subunits in the filament also plays an important role in its disassembly and turnover. When an ATP-actin monomer associates with the barbed end of the filament, its non-covalently bound ATP undergoes rapid hydrolysis (0.3 s−1), producing ADP-Pi-actin subunits. In a much slower step (0.007 s−1), the inorganic phosphate (Pi) dissociates, producing ADP-actin subunits. Actin filaments in these two different nucleotide states (ADP-Pi and ADP) have large differences in their kinetic properties, with ADP-actin subunits dissociating much faster than ADP-Pi-actin subunits from filament ends. In this manner, a filament “ages” due to its nucleotide state. At steady state, filaments undergo treadmilling, where there is net growth at the barbed ends and net depolymerization at the pointed ends. The rate-limiting step in treadmilling is dissociation of ADP-actin subunits from the pointed ends (0.27 s−1; ), and therefore, in the absence of additional cellular factors, filament turnover is relatively slow. However, actin structures in vivo often turnover at much faster rates (Fig. 1), as first revealed in the pioneering work of Yu-li Wang (Box 2). This now can be explained in large part by the activities of the actin disassembly-promoting factors highlighted in this review.

Our understanding of cellular actin turnover took a major leap forward with the pioneering work of Yu-li Wang, who used live cell imaging to directly visualize actin turnover at the leading edge for the first time. Prior to this work, the turnover of F-actin in cells was presumed by many to be slow, based on the biochemical properties of filaments assembled from purified actin, which turn over (or treadmill) very slowly. The rate-limiting step in this turnover is the dissociation of subunits from the pointed ends of filaments (0.27 s−1). At this rate, a filament that is 1 μm long requires ∼15 min to turn over. However, Wang microinjected cells with fluorescent actin and monitored fluorescence recovery after photobleaching at the leading edge, observing a turnover rate of 0.8 μm/min, at least 10-fold faster than the turnover of purified filaments. While this study was met with initial skepticism raised by concerns that photodamage might be leading to the unexpectedly fast turnover, subsequent studies from Theriot and Mitchison observed similarly high rates of turnover by photoactivation of actin instead of bleaching, both at the leading edge and in Listeria tails. This was later confirmed at the leading edge by powerful fluorescence speckle microscopy observations, directly visualizing the turnover of individual GFP-actin molecules. As live-imaging techniques have steadily improved and become widely used, actin turnover has been examined for a broad range of cellular actin arrays in different organisms. Collectively, this work has demonstrated that the turnover rates of cellular actin structures vary tremendously, ranging from seconds or minutes to hours or days (see Fig. 1).

For the purpose of discussing the mechanisms that drive actin network turnover, we have deconstructed this process into five discrete steps. However, it is important to keep in mind that each step can occur simultaneously within a network. The steps are: (1) curbing barbed end growth, (2) pruning branched filaments, (3) severing filaments with concomitant capping of new barbed ends, (4) promoting depolymerization from filament ends, and (5) recycling ADP-actin monomers back to ATP-actin monomers for new rounds of assembly. Each step is driven by one or more molecular mechanisms and is used to either (a) maintain a network’s size and shape while turning over its constituent filaments, which we refer to as the “dynamic maintenance” pathway, or (b) induce net disassembly of the network, which we refer to as the “demolition” pathway (Fig. 2). Networks that undergo dynamic maintenance include microvilli, stereocilia, sarcomeres, and stress fibers, where the size and shape of the actin structure remain fairly constant, while its individual filaments continue to undergo turnover (Fig. 1). This phenomenon requires balanced assembly and disassembly within the network, regardless of whether the turnover rate is slow or fast. Networks that undergo demolition include a retracting filopodium, a retracting lamellipodia, and a cortical endocytic patch disassembling after its relatively short lifetime (;;;;; Fig. 1).

One of the main purposes of this review is to summarize our current understanding of the conserved mechanisms regulating actin turnover and to highlight their molecular complexity. For many years, studies on cellular actin disassembly and turnover centered on the actions of only three proteins, actin depolymerizing factor (ADF)/cofilin, CP, and profilin. CP would terminate barbed end growth, ADF/cofilin would sever and depolymerize filaments, and profilin would recycle actin monomers for new rounds of assembly. However, it has become clear that these proteins are not sufficient to drive rapid actin turnover in vivo. Instead, at least five other ubiquitous factors are required: glia maturation factor (GMF), twinfilin, Srv2/cyclase-associated protein (CAP), coronin, and AIP1. These eight proteins constitute a core actin disassembly-promoting machinery in cells, while additional factors found in different organisms and cell types are undoubtedly involved. Importantly, the eight proteins we focus on here are conserved from fungi to animals and expressed in all known animal cell types. Further, each protein has distinct activities and functional role(s) in driving actin disassembly and/or recycling (Fig. 3). Cofilin, CP, AIP1, and Srv2/CAP are additionally found in plants. Further, some of these proteins are found in protists. For example, malaria-causing plasmodium expresses cofilin, profilin, coronin, CP, and a truncated version of CAP, and flagellated Leishmania additionally expresses twinfilin. Notably, three of the eight proteins (ADF/cofilin, GMF, and twinfilin) are structurally related to each other as members of the ADF-homology (ADF-H) domain family. Below, we discuss how specific combinations of these proteins collaborate through multicomponent mechanisms to drive distinct steps in actin network turnover.

Step 1: Curbing barbed end growth

In both the demolition and dynamic maintenance pathways, an important control point in network turnover is the attenuation of actin filament growth. This is achieved through proteins that cap the barbed ends of filaments (Fig. 2). Proteins that can either transiently or more stably cap barbed ends and block polymerization include CP, Eps8, adducin, gelsolin, twinfilin, villin, and IQGAP1. Among these proteins, the activity of CP is the best understood. CP is ubiquitously expressed, abundant in the cytosol (1–2 µM in yeast and mammalian cells;;; ), and arrests filament growth upon binding to the barbed end. In the absence of other cellular factors, CP remains stably bound to the barbed end, on average for tens of minutes. Through this activity, CP plays an important role in promoting the formation of actin networks comprised of short, branched filaments such as those found at the leading edge of motile cells and sites of endocytosis. Further, a recent study revealed that CP and the WH2 domains of Arp2/3 nucleation-promoting factors (NPFs) compete for filament barbed ends. This competition with CP liberates NPFs from barbed ends to bind actin monomers, indirectly stimulating Arp2/3-dependent branched actin nucleation. This mechanism may explain earlier observations showing that while CP limits the growth of individual filaments in networks, at the same time it promotes branched nucleation.

In addition to its roles in governing the formation of branched actin networks, CP has been shown to join formins at the barbed ends of actin filaments in vitro to form transient “decision complexes” (Fig. 4 A;; ). The formation of these complexes increases the rate of dissociation of both CP and formin from the barbed end by 10- to 50-fold. Moreover, the frequency of these transitions at the barbed end can be increased by ligands of CP (e.g., twinfilin) and formin (e.g., IQGAP1; Fig. 4 A;, Preprint; ). Collectively, these associative competition mechanisms promote rapid transitions between states of barbed end growth and capping in vitro. In vivo, this competitive interplay between CP and formins is important for maintaining the characteristic size, architecture, and dynamics of cellular actin networks. For instance, in yeast cells the deletion of CP leads to enlargement of the cortical actin patches nucleated by Arp2/3 complex, which results in aberrant recruitment of formins and tropomyosins to cortical patches, altering patch dynamics. Such interplay between CP and formins could also play an important role in filopodial extension. Filopodia grow by subunit addition at their distal tips, where they harbor formins and CP, which have been shown to have antagonistic roles in filopodia formation and dynamics. After filopodia grow to reach suitable lengths, growth pauses and filopodia abruptly switch to a state of rapid disassembly and shortening (Fig. 1;;; ). These acute transitions from growth to demolition may involve displacement of formins from the barbed ends by CP, IQGAP1, and/or other proteins.

Additional layers of barbed end regulation are achieved by other ligands of CP. This includes the CP inhibitor capping protein, Arp2/3, and myosin I linker (CARMIL), which localizes to the leading edge and strongly catalyzes CP displacement from barbed ends in vitro. Through this activity, CARMIL regulates lamellipodial protrusion and cell polarity. Another abundant inhibitor of CP is V-1/myotrophin, which does not displace CP from barbed ends, but instead sequesters CP in an inactive state in the cytoplasm, helping to restrict CP activity to the leading edge.

A third CP ligand, twinfilin (Fig. 3), has multiple effects on the interactions of CP with the barbed end. The C-terminal tail of twinfilin contains a CP-interacting (CPI) motif through which it binds directly to CP with high affinity and competes with other CPI proteins like CARMIL. Although CARMIL and twinfilin both displace CP from barbed ends, the uncapping activity of CARMIL is much stronger than that of twinfilin (180-fold versus sixfold faster dissociation of CP;; ). This difference, together with the observation that CARMIL is restricted to the plasma membrane while twinfilin is found throughout leading edge actin networks and in the cytosol, suggests that they may have spatially distinct roles in regulating CP. Additional insights into the twinfilin-CP mechanism have come from a crystal structure of twinfilin bound to CP and two actin subunits, and from a recent single molecule study showing that twinfilin molecules join CP at the barbed end in many repeated, short-lived interactions before CP displacement occurs.

In addition to its roles in leading edge actin networks, twinfilin localizes to endocytic actin patches and the tips of filopodia and stereocilia. Genetic observations in some of these studies have shown that twinfilin regulates the size and turnover of these actin-based structures, which may result from twinfilin functions at the barbed ends of filaments. Importantly, twinfilin not only promotes CP dissociation but also has important roles in driving depolymerization at filament ends, as described below (Fig. 4, A and D).

Together, these observations suggest that a large number of cellular factors dynamically compete for filament ends and/or interact with each other at filament ends, collectively attenuating barbed end growth.

Step 2: Pruning branched filaments

Following their assembly, the shape and filamentous architecture of actin networks can be rapidly transformed or “remodeled.” Such transitions are brought about through filament debranching, severing, and crosslinking mechanisms. In this section, we focus on debranching, which is specific to actin networks assembled by the branch-nucleating Arp2/3 complex. Initially, these actin structures are assembled as densely branched arrays, optimized for exerting force on membranes to drive protrusion and/or invagination. However, as the filaments within the networks age, they move inward from the cortex due to continued polymerization at the cortex and actomyosin retrograde flow. As the networks move in, filaments are pruned, i.e., daughter branches dissociate from mother filaments (see Fig. 3 in;;; ). In vivo, it is likely that debranching contributes not only to network remodeling but also to network disassembly, as it exposes (or uncaps) pointed ends of daughter filaments. In vitro, the filament branch junctions formed by Arp2/3 complex are extremely stable, persisting for tens of minutes, whereas in vivo they can turn over in 2–30 s. This disparity in rates in vitro versus in vivo suggests that cells have mechanisms for catalyzing debranching. To date, three proteins have been strongly implicated in promoting debranching: cofilin, coronin, and GMF.

Cofilin catalyzes debranching in addition to its more well-established roles in actin filament severing and depolymerization (;;; Fig. 4 B). Importantly, cofilin shows distinct concentration dependencies in its debranching and severing activities, suggesting that debranching occurs by a distinct mechanism. Recent total internal reflection fluorescence (TIRF) studies using labeled Arp2/3 complex verify that cofilin induces debranching (which removes Arp2/3 complex) rather than severing near branch junctions that would leave Arp2/3 complex on the filament side. The same study also showed that debranching events are preceded by the appearance of labeled cofilin somewhere close to the branch junction. Despite these advances, the molecular mechanism by which cofilin induces debranching remains poorly understood and merits further investigation.

A second protein implicated in debranching is coronin. Mammals have seven coronin genes, whereas yeast has only one (Fig. 4 B). Yeast coronin and several of the mammalian coronin isoforms bind Arp2/3 complex in addition to F-actin (Fig. 3;;;;;; ). Most coronin isoforms oligomerize and crosslink actin filaments in vitro. A role for coronin in debranching was first reported when depletion of mammalian coronin-1B was found to increase the density of branched filaments and barbed ends close to the leading edge and alter lamellipodial dynamics. Further, purified coronin-1B increased the in vitro debranching rate, although the underlying mechanism is still unclear. More recently, it was shown that coronin-7, which has a domain structure distinct from other coronin isoforms, also promotes debranching. It will be important to define the mechanism underlying these effects and determine whether debranching activities extend to coronin homologs in other model organisms.

The third protein implicated in debranching is GMF. GMF was first isolated from brain extracts and reported to be a cellular differentiation factor, but then was discovered to be a member of the ADF-H protein family and a component of the actin cytoskeleton involved in cell motility and endocytosis (Fig. 3;;; ). Biochemical analysis of yeast GMF revealed that it does not bind to actin, but instead has high affinity (Kd = 10 nM) for Arp2/3 complex (;; Fig. 4 B). Through this direct interaction, yeast and mammalian GMF isoforms induce debranching in vitro and suppress Wiskott Aldrich syndrome protein (WASP) and Arp2/3 complex–mediated branched actin nucleation. Consistent with these in vitro activities on Arp2/3 complex, overexpression of GMF in yeast leads to a reduction in the number of cortical actin patches and a concomitant increase in actin cables. Knockdown of Drosophila GMF disrupts lamellipodial retraction and causes defects in border cell migration. Further, Bear and colleagues utilized a point mutation in GMF (gmf1-101), which disrupts its debranching activity but not its ability to inhibit branched nucleation, to demonstrate that GMF’s debranching activity is required for normal Arp2/3 turnover rate at the leading edge and for proper lamellipodial protrusion and retraction dynamics.

How then does GMF destabilize the branch junction? GMF makes two important molecular contacts that induce debranching, one with Arp2/3 complex and one with the first actin subunit in the daughter filament. The Arp2/3-binding site on GMF occupies the same position in the ADF-H fold as the F/G-actin–binding site on cofilin, whereas the daughter filament-binding site on GMF spatially corresponds to the F-actin–binding site on cofilin (Fig. 3;; ). Thus, the mechanisms by which GMF induces debranching and cofilin induces severing appear to be structurally related. In contrast, the mechanism by which cofilin induces debranching is not well understood.

Another important step in debranching is ATP hydrolysis and phosphate release from the Arp2 and Arp3 subunits of the Arp2/3 complex. These changes in nucleotide state are set in motion by nucleation of the daughter filament and are required for proper actin network turnover in vivo. Whether coronin, cofilin, and/or GMF promote debranching in part by altering the rates of ATP hydrolysis and/or phosphate release on Arp2/3 complex needs to be investigated. Interestingly, GMF exhibits higher affinity for ADP- compared to ATP-bound Arp2/3 complex and more effectively destabilizes aged branches, so there is potential for multiple debranching factors working in concert to catalyze debranching. Finally, mechanical force can promote daughter filament detachment and possibly contribute to each of the biochemical debranching mechanisms above.

One of the next challenges is to determine how the activities of these three debranching factors (cofilin, coronin, and GMF) are coordinated in vivo and how their activities are counterbalanced by known branch stabilizers such as Abp1 and cortactin. Yeast coronin and GMF have distinct binding sites on the Arp2/3 complex and synergistically inhibit Arp2/3-mediated actin nucleation. Further, yeast GMF and cofilin have additive debranching effects, suggesting that they destabilize branches via distinct and complementary mechanisms. Yeast Abp1, which is another conserved ADF-H family protein, competes with GMF for Arp2/3 complex binding and antagonizes the debranching activity of GMF. While cortactin has long been known to stabilize branches against spontaneous debranching, it is unknown how it affects branch stability in the presence of branch destabilizers. Thus, there is much to learn about how branch stability is tuned in vivo through competitive interplay between branch destabilizers and stabilizers.

Step 3: Filament severing and capping

A central mechanism cells use to promote actin network disassembly is severing, which amplifies the number of filament ends (barbed and pointed) from which subunits can dissociate. However, because the cytosol contains high concentration of actin monomers, severing without concomitant barbed end capping can instead stimulate F-actin assembly. Indeed, severing by cofilin amplifies the number of barbed ends and in the presence of actin monomers promotes F-actin assembly in vitro and at the leading edge of cells. However, when severing is accompanied by rapid capping of the newly generated barbed ends it strongly increases the rate of F-actin disassembly. Barbed end cappers that may be involved in this process in vivo include CP, gelsolin, Eps8, twinfilin, WAVE1, adducins, villin, and IQGAP1.

Across the animal, plant, and fungal kingdoms, cofilin is the most abundant and well-characterized severing protein, although other proteins with severing activity may also be involved, e.g., gelsolin, villin, spire, cordon-bleu, and specific formins. Mammals have three distinct cofilin genes which encode cofilin-1, ADF, and cofilin-2. Cofilin-1 is ubiquitously expressed, ADF is found primarily in neuronal, epithelial, and endothelial tissues, and cofilin-2 is expressed in heart and skeletal muscle tissue. All three cofilins sever filaments and promote actin disassembly to different extents in vitro. In addition, all three cofilins promote depolymerization at filament ends in vitro (see next section). For the remainder of the review, we refer to cofilin proteins collectively as “cofilin” in order to focus on their shared properties; however, in a few instances we refer to their specific names to highlight differences.

Cofilins are essential for a wide range of actin-based processes that require rapid actin turnover, e.g., endocytosis, cell migration, cytokinesis, and cell morphogenesis. The mechanism by which cofilin severs filaments is well studied and has been reviewed elsewhere. In brief, cofilin binds preferentially to “aged” ADP-actin (monomeric and filamentous), but also catalyzes Pi release from ADP-Pi subunits in filaments, possibly via allosteric effects. Cofilin uses two distinct actin-binding surfaces (Fig. 3) to bind filaments cooperatively and dramatically alters F-actin conformation and mechanical properties. This leads to local discontinuities between the cofilin-decorated and “bare” regions on a filament, promoting breaks. TIRF microscopy and high-speed atomic force microscopy (AFM) studies further show that multiple cofilin molecules form a cluster on filament sides, which leads to conformational effects that propagate toward the pointed end. However, high-resolution electron microscopy indicates that the most obvious conformational changes in F-actin are restricted to the subunits where cofilin is bound. Thus, further structural work is needed to resolve whether cofilin perhaps has more subtle and longer-range allosteric effects on F-actin that influence filament dynamics and/or interactions of other actin-binding proteins. Cofilin severing also can be enhanced by torsional stresses imposed by filament cross-linkers and/or contractile forces induced by myosin motors.

On its own, cofilin has several limitations as a disassembly-promoting factor, and its activities change considerably in the presence of its cellular co-factors AIP1, Srv2/CAP, and coronin. The first limitation is that cofilin binds to F-actin slowly, with an on-rate of only ∼105 s−1 M−1. Second, filaments extensively decorated by cofilin alone do not sever. On its own, cofilin only severs filaments efficiently at nanomolar concentrations, where decoration is sparse. Third, as mentioned earlier, severing amplifies the number of barbed ends, and can therefore promote F-actin assembly rather than disassembly if the newly generated barbed ends are not rapidly capped. Importantly, each of these limitations is overcome by cofilin working with AIP1, coronin, and Srv2/CAP (Fig. 4 C). The structures of these three proteins and their actin-binding surfaces are highly conserved (Fig. 3), and their activities and mechanisms are summarized below. More information can also be found in reviews that focus on each protein.

Among the three co-factors, AIP1 is the only one known to directly bind cofilin. In vitro studies employing bulk and TIRF assays have shown that AIP1 works with cofilin to accelerate F-actin severing, and possibly pointed end depolymerization, both at the scale of single filaments and reconstituted actin networks. Further, filaments heavily decorated by cofilin are rapidly severed in the presence of AIP1. Thus, AIP1 and cofilin together overcome one of the above-mentioned limitations of cofilin alone. These in vitro effects are strongly supported by genetic interactions between cofilin and AIP1 and by the in vivo observation that loss of AIP1 leads to cofilin-decorated hyper-stabilized actin cables in yeast. Importantly, electron microscopy has revealed that in vitro AIP1 and cofilin together greatly reduce the size of F-actin severing products compared to cofilin alone. Whereas cofilin alone severs filaments into fragments ∼300–500 nm in length, AIP1 and cofilin together produce fragments only ∼50 nm long. Thus, AIP1 not only enhances the kinetics of severing, but also the extent of fragmentation.

Even more impressive than cofilin-AIP1 synergy is the combined activity of the coronin, cofilin, and AIP1 (or “CCA”) trio. Pioneering work from Brieher and colleagues identified these three proteins in crude mammalian cell extracts as factors that work together to drive rapid disassembly of Listeria actin tails even under assembly-promoting conditions. Subsequent single molecule analysis showed that the CCA mechanism proceeds by an ordered pathway (; Fig. 4 C). In the first step, coronin binds to F-actin. Next, coronin recruits cofilin to the same sites on the filament, accelerating cofilin accumulation on filament sides. Finally, AIP1 joins cofilin at these sites, and immediately induces severing. In this manner, the CCA mechanism severs filaments with much higher efficiency than cofilin alone. Moreover, all three proteins remain at or near the newly generated barbed ends and block new growth. High concentrations of AIP1 and cofilin together are also sufficient to suppress barbed end growth in vitro even in the absence of coronin, and this function is supported by genetic interactions between AIP1 and CP. However, further addition of coronin enhances these activities of cofilin and AIP1, leading to tight suppression of growth at newly generated barbed ends. CCA also has been reported to induce “catastrophic bursting” (the rapid disappearance of long stretches of F-actin from filament ends;; ). This potentially exciting mechanism merits further exploration, including determination of whether it involves mechanistic steps distinct from enhanced severing and depolymerization.

TIRF studies have shown that the three human cofilin isoforms have quantitative differences in their severing and depolymerization activities, yet all are enhanced by mammalian coronin and AIP1. Further, yeast coronin (Crn1) accelerates yeast cofilin recruitment to filament sides, and yeast AIP1 strongly enhances cofilin-mediated severing and disassembly. Therefore, CCA activities appear to be conserved. However, it is still unclear whether mammalian coronin isoforms besides coronin-1B participate in multi-component disassembly mechanisms, and how similar or different they are from each other. Another unresolved issue is the structural basis for CCA synergy. While progress has been made in determining the electron microscopy structures of F-actin decorated individually by cofilin or coronin, a deeper understanding will require high-resolution structures of filaments decorated by all three proteins. Dissection of such a complex mechanism may also benefit from the numerous separation-of-function mutants available in each of these proteins. While there are many details of the CCA mechanism yet to be resolved, it has served as an elegant illustration of how multiple cellular factors work together to produce biological effects that are not observed for any individual components.

A third co-factor that works with cofilin is CAP, which is called Srv2 in yeast. This 57-kD protein oligomerizes to form several distinct high-molecular-weight species seen by metal-shadowing electron microscopy. Subsequent negative-stain single particle averaging studies revealed that the N-terminal halves of yeast and mouse CAP (N-CAP) form hexameric shuriken structures with six actin-binding “blades,” each comprised of a helical-folded domain (HFD; Fig. 3;;;; ). However, more recent analyses of Xenopus and mouse CAP proteins, using analytical ultracentrifugation and AFM, suggest that CAP may form tetramers. Thus, CAP may be capable of adopting distinct oligomeric states, possibly with different activity states.

CAP has multiple domains and in vitro activities in promoting actin turnover (Fig. 4, C–E). The C-terminal half of CAP (C-CAP) dimerizes and recycles actin monomers, stimulating nucleotide exchange (ATP for ADP) for new rounds of actin assembly. N-CAP oligomerizes and binds autonomously to the sides of actin filaments to enhance cofilin-mediated severing by four- to eightfold, but has no severing effects on its own. The enhanced severing activity of N-CAP depends on a conserved F-actin–binding surface on its HFD (Fig. 3) and its oligomerization domain. Moreover, mutations at these sites cause in vivo defects in actin organization and endocytosis. Importantly, CAP does not interact directly with cofilin, and unlike coronin it does not enhance cofilin recruitment to filament sides. Instead, CAP reduces the time between cofilin binding and severing. Thus, CAP and coronin appear to have distinct, complementary roles in enhancing cofilin-mediated F-actin disassembly.

N-CAP also has strong effects in accelerating pointed end depolymerization, particularly in combination with cofilin (see next section for details). These effects also depend on the HFD. Kotila and colleagues showed that the HFD binds to surfaces on actin exposed at the pointed ends of filaments, which led them to call into question whether N-CAP is capable of binding to filament sides or enhancing cofilin-mediated severing. However, multiple lines of evidence have demonstrated that N-CAP binds to filament sides: (1) In co-sedimentation assays, N-CAP pellets with F-actin (Kd ∼2 µM; ). (2) In the same study, N-CAP was directly visualized on filament sides by EM, and induced “untwisting” of F-actin. (3) Further, N-Srv2 and N-CAP enhanced cofilin-mediated severing in three separate open-flow TIRF studies from two different groups. In contrast, Kotila and co-workers found that N-CAP failed to stimulate cofilin-mediated severing in microfluidic-TIRF experiments. However, flow-induced tension on filaments may have altered N-CAP binding. Thus, further investigation is needed to fully resolve whether CAP associates with filament sides and enhances cofilin-mediated severing, and how these interactions might be influenced by filament tension.

Another cellular factor that works with cofilin to promote actin network disassembly is the enzyme Molecules Interacting with CasL (MICAL; for details, see Box 3). Briefly, when MICAL binds to F-actin, it is activated, resulting in oxidation of two conserved methionine residues in the D-loop of actin (subdomain II). TIRF microscopy studies show that oxidation of actin by MICAL promotes severing as well as depolymerization at both ends of the filament, and that these activities are synergistic with those of cofilin.

An important but often overlooked mechanism in actin network turnover is destabilization of filaments by PTM of actin, induced by either cellular enzymes or bacterial toxins. The best-understood PTM that governs actin turnover is oxidation of Met44 in the D-loop of actin by the redox protein MICAL. Modification of even a small fraction of subunits in a filament leads to severing and/or depolymerization. Further, these effects are synergistic with cofilin-mediated severing in promoting rapid filament turnover in vitro and in vivo. The covalently modified actin monomers released by these mechanisms are compromised for assembly, but can be restored to an assembly-competent state by SelR, a methionine sulfoxide reductase enzyme that reverses oxidation of Met44 on actin. Thus, through activation and inactivation of MICAL and SelR, cells can rapidly and reversibly remodel their actin cytoskeletons in response to cues. These activities of MICAL are important for cytokinesis, axon guidance, and normal cellular actin dynamics. In addition, several bacterial pathogens induce disassembly of their host cell’s actin cytoskeleton by producing toxins that modify actin subunits through ADP-ribosylation or covalent crosslinking. Further, there are normal mechanisms in eukaryotic cells for introducing similar modifications, e.g., transglutaminases, which crosslink actin subunits, and transferase A, which ADP-ribosylates actin. Crosslinking of actin monomers leads to an accumulation of polymerization-deficient dimers, trimers, and oligomers, depleting the monomer pool and causing the disassembly of F-actin structures. Depending on the specific residue modified, ADP-ribosylation of actin differentially affects actin polymerization. For instance, ADP-ribosylation of Arg177 compromises polymerization and blocks barbed end growth, lowering F-actin levels, whereas ADP-ribosylation of Thr148 leads to higher F-actin levels, possibly by weakening thymosin-β4 interactions with the modified actin monomers. Additional modifications of actin that can occur in vivo and can influence actin turnover include arginylation, acetylation, phosphorylation, methylation, and nitrosylation. Collectively, these PTMs are opening new avenues for actin regulation research and ultimately must be considered alongside the multicomponent disassembly mechanisms described herein.

Collectively, the examples discussed above emphasize how cells employ multi-component molecular mechanisms to promote the rapid severing, capping, and depolymerization of actin filaments.

Step 4: Accelerated depolymerization

In both the demolition and dynamic maintenance pathways, the depolymerization of filaments is a critical step in network turnover (Fig. 2). In the demolition pathway, the F-actin fragments produced by severing must be depolymerized into monomers, so that they can be recycled for use in new rounds of actin assembly. In the dynamic maintenance pathway, actin arrays of a specified length and architecture (e.g., microvilli or stereocilia) are maintained dynamically, due to balanced polymerization at one end of the network and depolymerization at the other end.

In the absence of cellular factors, the rate of subunit dissociation from pointed ends is slow (0.27 s−1; ), and at this rate even the shortest fragments generated by cofilin and AIP1 severing (∼20 subunits long;; ) would still require over 1 min to depolymerize. Such slow rates are incompatible with the fast speeds of actin turnover observed in vivo for many actin networks (Fig. 1), suggesting that cells are likely to have mechanisms for accelerating subunit dissociation from filament ends. However, this concept was vigorously debated for many years, due primarily to a lack of direct observation of such effects. However, the debate was finally settled by a series of in vitro TIRF microscopy studies in 2015–2019, which directly visualized accelerated depolymerization at filament ends induced by twinfilin, cofilin, and/or Srv2/CAP (Fig. 4 D;;;;;,; ).

The first of these studies showed that yeast Srv2/CAP and twinfilin jointly accelerate filament depolymerization at barbed ends (by approximately threefold) and pointed ends (by ∼20-fold; ). Twinfilin is a conserved member of the ADF-H domain protein family, with an unusual molecular architecture consisting of two ADF-H domains separated by a short linker, followed by a C-terminal tail that interacts with capping protein (Fig. 3;;; ). Twinfilin binds with high affinity to ADP-actin monomers and very low affinity to filament sides. Early genetic interactions between twinfilin and cofilin mutants had suggested a role for twinfilin in promoting actin turnover; however, for many years the only known activities of twinfilin were actin monomer sequestering and a poorly understood barbed end capping effect. TIRF studies then revealed that yeast twinfilin accelerates depolymerization at barbed ends, and together with yeast Srv2/CAP strongly accelerates depolymerization at pointed ends. Single molecule imaging further showed that twinfilin directly associates with barbed ends (Fig. 4 D).

In mammals, the three twinfilin isoforms (Twf1, Twf2a, and Twf2b) all collaborate with mammalian CAP1 to accelerate barbed end depolymerization, similar to their yeast counterparts. However, unlike the yeast proteins, mammalian twinfilins and CAP only modestly accelerate pointed end depolymerization. While there is some evidence to suggest that twinfilin interacts processively at barbed ends, these data are far from conclusive, and there have been other observations pointing instead to a non-processive mechanism in which each twinfilin molecule removes only one to two subunits per binding event at the barbed end. The effects of twinfilin on barbed end depolymerization also depend on the nucleotide state (or “age”) of the filament, as twinfilin accelerates the depolymerization of ADP-Pi, but slows the depolymerization of ADP F-actin. Further, these activities are observed even under assembly-promoting conditions, i.e., in the presence of actin monomers. Thus, twinfilin appears to be a specialized depolymerase, capable of bypassing the normal filament aging step in disassembly in vitro. It is possible that these unique activities of twinfilin are employed in vivo to accelerate the disassembly of newly polymerized regions of cellular actin networks.

The ability of cofilin to accelerate filament depolymerization was validated in 2017 by two groups employing in vitro microfluidics-assisted TIRF microscopy. Earlier evidence from in vitro bulk assays had argued that cofilin increases the rate of subunit dissociation from pointed ends. However, in the absence of direct visualization of these effects, this model remained controversial, especially since severing had been observed in EM and TIRF studies. However, the studies above ended this debate by directly observing barbed and pointed ends of filaments shortening at an accelerated rate in the presence of all three mammalian cofilin isoforms. Interestingly, once filaments became heavily decorated along their sides by cofilin, only the pointed ends continued to depolymerize faster than control filaments, whereas the barbed ends depolymerized more slowly than control filaments. Furthermore, under these conditions (saturating cofilin) the barbed ends depolymerized persistently, even in the presence of actin monomers or capping protein. Differences in the depolymerization rates were observed for cofilin isoforms. ADF accelerated depolymerization by as much as 22-fold at pointed ends (Fig. 4 D), whereas cofilin-1 and cofilin-2 had only two- to threefold effects on depolymerization at pointed ends. These differences between ADF and cofilin-1 could possibly explain why many non-muscle cell types express both isoforms.

In 2019, two groups used microfluidics-assisted TIRF to show that Srv2/CAP synergizes with cofilin in depolymerizing the pointed ends of filaments. These studies showed that N-CAP alone accelerates pointed end depolymerization by approximately fivefold, but together with cofilin induces depolymerization >300-fold faster than control filaments, reaching rates of ∼50 subunits s−1. These striking effects were observed at concentrations of CAP and cofilin similar to their cellular concentrations. Further, the mechanism involved CAP molecules processively tracking the pointed ends of cofilin-decorated filaments (Fig. 4 D). The pointed end depolymerization effects induced by yeast twinfilin and CAP may involve a related mechanism. Consistent with this idea, mutations in the conserved actin-binding surface of the HFD domain of CAP abolish its in vitro ability to jointly catalyze pointed end depolymerization with either twinfilin or cofilin. The crystal structure of a ternary HFD-actin-cofilin complex further suggests that the pointed end depolymerization effects of CAP may require interactions of HFD domains with both actin subunits exposed at the pointed end. In vivo, the robust pointed end depolymerization mechanism of CAP and cofilin could be used to rapidly depolymerize the F-actin fragments resulting from severing and/or to maintain rapid turnover at one end of a cellular actin network decorated by tropomyosin, which protects filament sides from severing. Indeed, mutations in the HFD that disrupt CAP depolymerization activities cause striking defects in actin organization and dynamics in vivo.

Step 5: Actin monomer recycling

Once ADP-actin monomers are released from filament ends (catalyzed by severing and depolymerization mechanisms), they must be recycled back to the ATP-bound state to be used for new rounds of filament assembly. Released ADP-actin monomers that are free, i.e., not bound to other factors, spontaneously undergo fast nucleotide exchange (ATP for ADP). However, due to the high affinities of cofilin and twinfilin for ADP-actin monomers, a substantial fraction of released ADP-actin monomers in the cytosol is predicted to be bound to cofilin and twinfilin. Importantly, cofilin and twinfilin also strongly suppress nucleotide exchange on actin monomers, as does thymosin β4, which is a highly abundant peptide in the cytosol of animal cells. How then, under these cytosolic conditions, are actin monomers recycled?

Two cellular factors, profilin and Srv2/CAP, have been implicated in overcoming these barriers to promote nucleotide exchange and recharging of monomers (Fig. 4 E;;;;; ). Both proteins are abundant and conserved throughout the plant, animal, and fungal kingdoms. Srv2/CAP is more effective than profilin in promoting nucleotide exchange on cofilin-bound ADP-actin monomers in vitro. This can be explained by the low affinity of profilin for ADP-actin monomers. In contrast, Srv2/CAP exhibits a high affinity for ADP-actin monomers (Kd ∼20 nM), and catalyzes cofilin dissociation from monomers. These activities of Srv2/CAP are mediated by its C-CAP, consisting of actin-binding WH2 and β-sheet/CAP and X-linked retinitis pigmentosa 2 protein (CARP) domains (Fig. 3;;;; ). A cocrystal structure of the C-CAP dimer bound to two actin monomers revealed the structural basis for CAP-accelerated nucleotide exchange on actin. Interestingly, the C-terminal tail of C-CAP (last four residues) directly binds to the nucleotide-sensing region of G-actin. Deletion of these residues abolishes C-CAP effects on nucleotide exchange in vitro and leads to pronounced defects in yeast cell growth and actin organization. C-CAP also contains a profilin-binding polyproline tract (P1), which binds directly to profilin. This interaction may facilitate Srv2/CAP’s “middleman” role in converting cofilin- and twinfilin-bound ADP-actin monomers into profilin-bound ATP-actin monomers ready for new rounds of assembly (Fig. 4 E;; ). Consistent with this idea, a recent study determined that CAP is the limiting factor for monomer recycling and sustained motility in a “closed” in vitro reconstitution system containing purified proteins (verproline central acidic on beads, plus actin, profilin, CP, Arp2/3 complex, cofilin, and CAP; ).

One important question for the future is how the actin monomer recycling functions of Srv2/CAP are integrated with its F-actin severing and depolymerization functions (discussed in the previous section). The filament severing and depolymerization activities of Srv2/CAP are mediated by its N-CAP, whereas monomer recycling is mediated by its C-CAP. Interestingly, the two halves of Srv2/CAP can function in trans as independent fragments in vitro and in vivo. Thus, the F-actin and G-actin turnover functions of Srv2/CAP appear to be fairly autonomous. This raises the intriguing question of why these two functions are linked in one molecule and whether physically joining the two halves of Srv2/CAP provides intramolecular regulation between these two functions and/or new as-yet-to-be-discovered activities.

Another key question is how the monomer pool is controlled in vivo. Cells maintain a large fraction of their total actin in a monomeric state (∼100 µM) so that high concentrations of monomers are available to drive rapid polymerization. In vertebrate cells, the largest fraction of the actin monomer pool is bound to thymosin-β4, owing to its high abundance. Thymosin-β4 blocks actin monomer addition to filament ends, i.e., is a sequestering protein. Importantly, thymosin-β4 interactions with monomers are highly transient and thus actin monomers can be rapidly handed off to profilin, which is not a sequestering protein. Profilin-bound monomers can be added to free barbed ends or used by formins and Ena/vasodilator-stimulated phosphoprotein to accelerate filament elongation. Therefore, due to the rapid exchange of actin monomers between thymosin-β4 and profilin, cells are able to maintain a homeostatic balance between their large pool of thymosin-β4–sequestered monomers (80–90 µM) and smaller pool of assembly-ready profilin-bound monomers (10–20 µM; Fig. 4 E;; ).

Deployment of actin turnover mechanisms in vivo

The mechanisms discussed above provide cells with a diverse toolkit for reshaping actin network architecture and tuning network turnover rates. For instance, branched networks at the leading edge of cells and sites of endocytosis start out as densely arborized arrays, but as the networks move inward, they are pruned (by debranchers like cofilin, coronin, and GMF), transforming their filamentous organization and properties. At the same time, some of these networks undergo severing and depolymerization, reducing average filament length. More work is needed to determine how network architecture is dynamically transformed in vivo. Indeed, a recent study showed that CAP, enhanced by its interactions with Abp1, dynamically slides and bundles actin filaments (branched and unbranched), coalescing them into compact bundles through a novel activity. This study also showed that specific mutations in CAP that disrupt these activities cause strong defects in actin organization in vivo. This is likely to be only one of many remodeling activities used in cells to dynamically reorganize actin networks.

How different turnover mechanisms are used in vivo may also depend on whether a network is earmarked for the dynamic maintenance or the demolition pathway (Fig. 2). Dynamic maintenance requires balanced assembly and disassembly at the two ends of the network, regardless of whether the turnover rate is slow or fast. However, we still do not understand how this balance is achieved. On the other hand, the demolition pathway requires strong inhibition of network assembly (Fig. 2). This may be achieved by locally shutting off actin nucleators and polymerases and by activating barbed end cappers and depolymerizers, coupled with filament severing and depolymerization. There is some evidence for this in filopodial retraction, as cofilin and twinfilin appear at filopodial tips just before they retract. Similarly, lamellipodial retraction is accompanied by the appearance of many disassembly-promoting factors, including cofilin, AIP1, coronin, Srv2/CAP, GMF, and twinfilin.

Another future challenge is to determine how the different actin-disassembly mechanisms are influenced by the presence of other actin-binding proteins decorating networks. For instance, tropomyosins (TPMs) are an abundant family of coiled-coil proteins that polymerize along the sides of filaments, decorating most known cellular actin structures. Mammalian cells express >40 different TPM isoforms generated by alternative splicing of four distinct genes. These isoforms exhibit a wide range of antagonistic effects on cofilin in vitro, which is thought to provide cells with the plasticity to differentially protect various actin structures. Indeed, specific roles for TPM isoforms are suggested by their distinct in vivo localization patterns to actin networks with different turnover dynamics and lifetimes. Cofilin can be antagonized in vitro by other members of the ADF-H protein family (e.g., Abp1, drebrin, and coactosin), which compete with cofilin for F-actin–side binding. This view is supported by genetic observations showing that loss of drebrin or coactosin leads to increased cofilin-mediated F-actin turnover. Mechanisms that accelerate pointed end depolymerization are also likely to be influenced by actin-binding proteins that cap and/or polymerize pointed ends, such as tropomodulins and VopF. Thus, there is still much to learn about how these and other side-binding proteins affect the multicomponent disassembly mechanisms discussed above.

Finally, it remains to be determined how actin network remodeling and turnover are differentially affected by the crosslinking proteins that spatially organize filaments into distinct higher-order networks. For instance, fascin and fimbrin bundle filaments into tightly packed linear arrays, e.g., in filopodia. Some studies suggest that fascin bundling enhances severing by cofilin, while other studies suggest that fascin blocks cofilin severing (, Preprint). Other longer-lived cellular actin structures are organized by a different set of crosslinkers (e.g., α-actinin, filamin, espin, calponin, and Eps8), which may endow networks with different degrees of resistance or vulnerability to specific disassembly mechanisms. Ultimately, understanding how various cellular actin networks (e.g., lamellipodia, filopodia, microvilli, stereocilia, and actin cables) are differentially remodeled and turned over will require a deeper knowledge of how their filament side-binding and crosslinking proteins influence the disassembly mechanisms, combined with how the activities of disassembly factors are locally regulated in the vicinity of each network.

Concluding remarks

Over the past two decades, our understanding of how cellular actin networks are disassembled has expanded profoundly due to a large body of work defining the structures, in vitro mechanisms, and biological roles of the disassembly and turnover machinery. One of the most important insights made is that distinct sets of disassembly factors work together in multicomponent mechanisms to produce emergent effects, as discussed above. These advances notwithstanding, more work is needed to define the structural basis of each mechanism and to determine which mechanisms are used to drive the turnover of which actin networks in vivo. Further, we only have a limited understanding of how these turnover mechanisms are regulated by signaling pathways through posttranslational modification (PTM) of actin and actin-binding proteins. Ultimately, gaining a more complete understanding of cellular actin network disassembly will require developing and combining diverse approaches, including high-resolution cell imaging, structural biology, in vitro multiwavelength single molecule imaging, optogenetics, theory, and modeling.

References 258

  1. 1.

    Akin, O., and R.D. Mullins. 2008. Capping protein increases the rate of actin-based motility by promoting filament nucleation by the Arp2/3 complex. Cell. 133:841–851.

    Not in our library10.1016/j.cell.2008.04.011

  2. 2.

    Aktories, K., A.E. Lang, C. Schwan, and H.G. Mannherz. 2011. Actin as target for modification by bacterial protein toxins. FEBS J. 278:4526–4543.

    Not in our library10.1111/j.1742-4658.2011.08113.x

  3. 3.

    Almenar-Queralt, A., A. Lee, C.A. Conley, L. Ribas de Pouplana, and V.M. Fowler. 1999. Identification of a novel tropomodulin isoform, skeletal tropomodulin, that caps actin filament pointed ends in fast skeletal muscle. J. Biol. Chem. 274:28466–28475.

    Not in our library10.1074/jbc.274.40.28466

  4. 4.

    Andrianantoandro, E., and T.D. Pollard. 2006. Mechanism of actin filament turnover by severing and nucleation at different concentrations of ADF/cofilin. Mol. Cell. 24:13–23.

    Not in our library10.1016/j.molcel.2006.08.006

  5. 5.

    Balcer, H.I., A.L. Goodman, A.A. Rodal, E. Smith, J. Kugler, J.E. Heuser, and B.L. Goode. 2003. Coordinated regulation of actin filament turnover by a high-molecular-weight Srv2/CAP complex, cofilin, profilin, and Aip1. Curr. Biol. 13:2159–2169.

    Not in our library10.1016/j.cub.2003.11.051

  6. 6.

    Berro, J., and T.D. Pollard. 2014. Synergies between Aip1p and capping protein subunits (Acp1p and Acp2p) in clathrin-mediated endocytosis and cell polarization in fission yeast. Mol. Biol. Cell. 25:3515–3527.

    Not in our library10.1091/mbc.e13-01-0005

  7. 7.

    Bertling, E., P. Hotulainen, P.K. Mattila, T. Matilainen, M. Salminen, and P. Lappalainen. 2004. Cyclase-associated protein 1 (CAP1) promotes cofilin-induced actin dynamics in mammalian nonmuscle cells. Mol. Biol. Cell. 15:2324–2334.

    Not in our library10.1091/mbc.e04-01-0048

  8. 8.

    Bertling, E., O. Quintero-Monzon, P.K. Mattila, B.L. Goode, and P. Lappalainen. 2007. Mechanism and biological role of profilin-Srv2/CAP interaction. J. Cell Sci. 120:1225–1234.

    Not in our library10.1242/jcs.000158

  9. 9.

    Bhattacharya, N., S. Ghosh, D. Sept, and J.A. Cooper. 2006. Binding of myotrophin/V-1 to actin-capping protein: Implications for how capping protein binds to the filament barbed end. J. Biol. Chem. 281:31021–31030.

    Not in our library10.1074/jbc.m606278200

  10. 10.

    Bibeau, J.P., N.G. Pandit, S. Gray, N. Shatery Nejad, C.V. Sindelar, W. Cao, and E.M. De La Cruz. 2023. Twist response of actin filaments. Proc. Natl. Acad. Sci. USA. 120:e2208536120.

    Not in our library10.1073/pnas.2208536120

  11. 11.

    Billault-Chaumartin, I., and S.G. Martin. 2019. Capping protein insulates Arp2/3-assembled actin patches from formins. Curr. Biol. 29:3165–3176.e6.

    Not in our library10.1016/j.cub.2019.07.088

  12. 12.

    Blanchoin, L., and T.D. Pollard. 1999. Mechanism of interaction of Acanthamoeba actophorin (ADF/Cofilin) with actin filaments. J. Biol. Chem. 274:15538–15546.

    Not in our library10.1074/jbc.274.22.15538

  13. 13.

    Blanchoin, L., and T.D. Pollard. 2002. Hydrolysis of ATP by polymerized actin depends on the bound divalent cation but not profilin. Biochemistry. 41:597–602.

    Not in our library10.1021/bi011214b

  14. 14.

    Blanchoin, L., T.D. Pollard, and R.D. Mullins. 2000. Interactions of ADF/cofilin, Arp2/3 complex, capping protein and profilin in remodeling of branched actin filament networks. Curr. Biol. 10:1273–1282.

    Not in our library10.1016/s0960-9822(00)00749-1

  15. 15.

    Blikstad, I., F. Markey, L. Carlsson, T. Persson, and U. Lindberg. 1978. Selective assay of monomeric and filamentous actin in cell extracts, using inhibition of deoxyribonuclease I. Cell. 15:935–943.

    Not in our library10.1016/0092-8674(78)90277-5

  16. 16.

    Boczkowska, M., G. Rebowski, and R. Dominguez. 2013. Glia maturation factor (GMF) interacts with Arp2/3 complex in a nucleotide state-dependent manner. J. Biol. Chem. 288:25683–25688.

    Not in our library10.1074/jbc.c113.493338

  17. 17.

    Bombardier, J.P., J.A. Eskin, R. Jaiswal, I.R. Corrêa Jr., M.Q. Xu, B.L. Goode, and J. Gelles. 2015. Single-molecule visualization of a formin-capping protein ‘decision complex’ at the actin filament barbed end. Nat. Commun. 6:8707.

    Not in our library10.1038/ncomms9707

  18. 18.

    Bonder, E.M., D.J. Fishkind, and M.S. Mooseker. 1983. Direct measurement of critical concentrations and assembly rate constants at the two ends of an actin filament. Cell. 34:491–501.

    Not in our library10.1016/0092-8674(83)90382-3

  19. 19.

    Bosch, M., K.H.D. Le, B. Bugyi, J.J. Correia, L. Renault, and M.F. Carlier. 2007. Analysis of the function of Spire in actin assembly and its synergy with formin and profilin. Mol. Cell. 28:555–568.

    Not in our library10.1016/j.molcel.2007.09.018

  20. 20.

    Bravo-Cordero, J.J., M.A.O. Magalhaes, R.J. Eddy, L. Hodgson, and J. Condeelis. 2013. Functions of cofilin in cell locomotion and invasion. Nat. Rev. Mol. Cell Biol. 14:405–415.

    Not in our library10.1038/nrm3609

  21. 21.

    Breitsprecher, D., S.A. Koestler, I. Chizhov, M. Nemethova, J. Mueller, B.L. Goode, J.V. Small, K. Rottner, and J. Faix. 2011. Cofilin cooperates with fascin to disassemble filopodial actin filaments. J. Cell Sci. 124:3305–3318.

    Not in our library10.1242/jcs.086934

  22. 22.

    Brieher, W.M., H.Y. Kueh, B.A. Ballif, and T.J. Mitchison. 2006. Rapid actin monomer-insensitive depolymerization of Listeria actin comet tails by cofilin, coronin, and Aip1. J. Cell Biol. 175:315–324.

    Not in our library10.1083/jcb.200603149

  23. 23.

    Cai, L., N. Holoweckyj, M.D. Schaller, and J.E. Bear. 2005. Phosphorylation of coronin 1B by protein kinase C regulates interaction with Arp2/3 and cell motility. J. Biol. Chem. 280:31913–31923.

    Not in our library10.1074/jbc.m504146200

  24. 24.

    Cai, L., A.M. Makhov, D.A. Schafer, and J.E. Bear. 2008. Coronin 1B antagonizes cortactin and remodels Arp2/3-containing actin branches in lamellipodia. Cell. 134:828–842.

    Not in our library10.1016/j.cell.2008.06.054

  25. 25.

    Cai, L., T.W. Marshall, A.C. Uetrecht, D.A. Schafer, and J.E. Bear. 2007. Coronin 1B coordinates Arp2/3 complex and cofilin activities at the leading edge. Cell. 128:915–929.

    Not in our library10.1016/j.cell.2007.01.031

  26. 26.

    Carlier, M.F., V. Laurent, J. Santolini, R. Melki, D. Didry, G.X. Xia, Y. Hong, N.H. Chua, and D. Pantaloni. 1997. Actin depolymerizing factor (ADF/cofilin) enhances the rate of filament turnover: Implication in actin-based motility. J. Cell Biol. 136:1307–1322.

    Not in our library10.1083/jcb.136.6.1307

  27. 27.

    Carlier, M.F., and D. Pantaloni. 1986. Direct evidence for ADP-Pi-F-actin as the major intermediate in ATP-actin polymerization. Rate of dissociation of Pi from actin filaments. Biochemistry. 25:7789–7792.

    Not in our library10.1021/bi00372a001

  28. 28.

    Carlier, M.F., D. Pantaloni, and E.D. Korn. 1987. The mechanisms of ATP hydrolysis accompanying the polymerization of Mg-actin and Ca-actin. J. Biol. Chem. 262:3052–3059.

    Not in our library10.1016/s0021-9258(18)61467-x

  29. 29.

    Carlier, M.F., and S. Shekhar. 2017. Global treadmilling coordinates actin turnover and controls the size of actin networks. Nat. Rev. Mol. Cell Biol. 18:389–401.

    Not in our library10.1038/nrm.2016.172

  30. 30.

    Carman, P.J., K.R. Barrie, G. Rebowski, and R. Dominguez. 2023. Structures of the free and capped ends of the actin filament. Science. 380:1287–1292.

    Not in our library10.1126/science.adg6812

  31. 31.

    Chan, A.Y., M. Bailly, N. Zebda, J.E. Segall, and J.S. Condeelis. 2000. Role of cofilin in epidermal growth factor-stimulated actin polymerization and lamellipod protrusion. J. Cell Biol. 148:531–542.

    Not in our library10.1083/jcb.148.3.531

  32. 32.

    Chan, C., C.C. Beltzner, and T.D. Pollard. 2009. Cofilin dissociates Arp2/3 complex and branches from actin filaments. Curr. Biol. 19:537–545.

    Not in our library10.1016/j.cub.2009.02.060

  33. 33.

    Chan, K.T., S.J. Creed, and J.E. Bear. 2011. Unraveling the enigma: Progress towards understanding the coronin family of actin regulators. Trends Cell Biol. 21:481–488.

    Not in our library10.1016/j.tcb.2011.04.004

  34. 34.

    Chaudhry, F., D. Breitsprecher, K. Little, G. Sharov, O. Sokolova, and B.L. Goode. 2013. Srv2/cyclase-associated protein forms hexameric shurikens that directly catalyze actin filament severing by cofilin. Mol. Biol. Cell. 24:31–41.

    Not in our library10.1091/mbc.e12-08-0589

  35. 35.

    Chaudhry, F., C. Guérin, M. von Witsch, L. Blanchoin, and C.J. Staiger. 2007. Identification of Arabidopsis cyclase-associated protein 1 as the first nucleotide exchange factor for plant actin. Mol. Biol. Cell. 18:3002–3014.

    Not in our library10.1091/mbc.e06-11-1041

  36. 36.

    Chaudhry, F., S. Jansen, K. Little, C. Suarez, R. Boujemaa-Paterski, L. Blanchoin, and B.L. Goode. 2014. Autonomous and in trans functions for the two halves of Srv2/CAP in promoting actin turnover. Cytoskeleton. 71:351–360.

    Not in our library10.1002/cm.21170

  37. 37.

    Chaudhry, F., K. Little, L. Talarico, O. Quintero-Monzon, and B.L. Goode. 2010. A central role for the WH2 domain of Srv2/CAP in recharging actin monomers to drive actin turnover in vitro and in vivo. Cytoskeleton. 67:120–133.

    Not in our library10.1002/cm.20429

  38. 38.

    Chen, J., D. Godt, K. Gunsalus, I. Kiss, M. Goldberg, and F.A. Laski. 2001. Cofilin/ADF is required for cell motility during Drosophila ovary development and oogenesis. Nat. Cell Biol. 3:204–209.

    Not in our library10.1038/35055120

  39. 39.

    Chen, Q., N. Courtemanche, and T.D. Pollard. 2015. Aip1 promotes actin filament severing by cofilin and regulates constriction of the cytokinetic contractile ring. J. Biol. Chem. 290:2289–2300.

    Not in our library10.1074/jbc.m114.612978

  40. 40.

    Chhabra, E.S., and H.N. Higgs. 2006. 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.

    Not in our library10.1074/jbc.m604666200

  41. 41.

    Chikireddy, J., L. Lengagne, R.L. Borgne, H. Wioland, G. Romet-Lemonne, and A. Jégou. 2023. Fascin-induced bundling protects actin filaments from disassembly by cofilin. bioRxiv. (Preprint posted May 19, 2023).

    Not in our library10.1101/2023.05.19.541460

  42. 42.

    Chin, S.M., S. Jansen, and B.L. Goode. 2016. TIRF microscopy analysis of human Cof1, Cof2, and ADF effects on actin filament severing and turnover. J. Mol. Biol. 428:1604–1616.

    Not in our library10.1016/j.jmb.2016.03.006

  43. 43.
    Competition between Tropomyosin, Fimbrin, and ADF/Cofilin drives their sorting to distinct actin filament networks · 2017
    Not checked
  44. 44.

    Chung, J., B.L. Goode, and J. Gelles. 2022. Single-molecule analysis of actin filament debranching by cofilin and GMF. Proc. Natl. Acad. Sci. USA. 119:e2115129119.

    Not in our library10.1073/pnas.2115129119

  45. 45.

    Clark, M.G., J. Teply, B.K. Haarer, S.C. Viggiano, D. Sept, and D.C. Amberg. 2006. A genetic dissection of Aip1p’s interactions leads to a model for Aip1p-cofilin cooperative activities. Mol. Biol. Cell. 17:1971–1984.

    Not in our library10.1091/mbc.e05-10-0956

  46. 46.

    Colin, A., T. Kotila, C. Guérin, M. Orhant-Prioux, B. Vianay, A. Mogilner, P. Lappalainen, M. Théry, and L. Blanchoin. 2023. Recycling of the actin monomer pool limits the lifetime of network turnover. EMBO J. 42:e112717.

    Not in our library10.15252/embj.2022112717

  47. 47.

    Collins, A., A. Warrington, K.A. Taylor, and T. Svitkina. 2011. Structural organization of the actin cytoskeleton at sites of clathrin-mediated endocytosis. Curr. Biol. 21:1167–1175.

    Not in our library10.1016/j.cub.2011.05.048

  48. 48.

    Del Duca, S., D. Serafini-Fracassini, P. Bonner, M. Cresti, and G. Cai. 2009. Effects of post-translational modifications catalysed by pollen transglutaminase on the functional properties of microtubules and actin filaments. Biochem. J. 418:651–664.

    Not in our library10.1042/bj20081781

  49. 49.

    DesMarais, V., I. Ichetovkin, J. Condeelis, and S.E. Hitchcock-DeGregori. 2002. Spatial regulation of actin dynamics: A tropomyosin-free, actin-rich compartment at the leading edge. J. Cell Sci. 115:4649–4660.

    Not in our library10.1242/jcs.00147

  50. 50.

    Devineni, N., L.S. Minamide, M. Niu, D. Safer, R. Verma, J.R. Bamburg, and V.T. Nachmias. 1999. A quantitative analysis of G-actin binding proteins and the G-actin pool in developing chick brain. Brain Res. 823:129–140.

    Not in our library10.1016/s0006-8993(99)01147-6

  51. 51.

    Dolge, L., K. Aufenvenne, H. Traupe, and W. Baumgartner. 2012. Beta-actin is a target for transglutaminase activity at synaptic endings in chicken telencephalic cell cultures. J. Mol. Neurosci. 46:410–419.

    Not in our library10.1007/s12031-011-9601-8

  52. 52.

    Edwards, M., Y. Liang, T. Kim, and J.A. Cooper. 2013. Physiological role of the interaction between CARMIL1 and capping protein. Mol. Biol. Cell. 24:3047–3055.

    Not in our library10.1091/mbc.e13-05-0270

  53. 53.

    Edwards, M., A. Zwolak, D.A. Schafer, D. Sept, R. Dominguez, and J.A. Cooper. 2014. Capping protein regulators fine-tune actin assembly dynamics. Nat. Rev. Mol. Cell Biol. 15:677–689.

    Not in our library10.1038/nrm3869

  54. 54.

    Elam, W.A., H. Kang, and E.M. De la Cruz. 2013a. Biophysics of actin filament severing by cofilin. FEBS Lett. 587:1215–1219.

    Not in our library10.1016/j.febslet.2013.01.062

  55. 55.

    Elam, W.A., H. Kang, and E.M. De La Cruz. 2013b. Competitive displacement of cofilin can promote actin filament severing. Biochem. Biophys. Res. Commun. 438:728–731.

    Not in our library10.1016/j.bbrc.2013.07.109

  56. 56.

    Falck, S., V.O. Paavilainen, M.A. Wear, J.G. Grossmann, J.A. Cooper, and P. Lappalainen. 2004. Biological role and structural mechanism of twinfilin-capping protein interaction. EMBO J. 23:3010–3019.

    Not in our library10.1038/sj.emboj.7600310

  57. 57.

    Föger, N., L. Rangell, D.M. Danilenko, and A.C. Chan. 2006. Requirement for coronin 1 in T lymphocyte trafficking and cellular homeostasis. Science. 313:839–842.

    Not in our library10.1126/science.1130563

  58. 58.
    Oxidation of F-actin controls the terminal steps of cytokinesis · 2017
    No notice
  59. 59.

    Fujiwara, I., K. Remmert, and J.A. Hammer III. 2010. Direct observation of the uncapping of capping protein-capped actin filaments by CARMIL homology domain 3. J. Biol. Chem. 285:2707–2720.

    Not in our library10.1074/jbc.m109.031203

  60. 60.

    Fujiwara, I., K. Remmert, G. Piszczek, and J.A. Hammer. 2014. Capping protein regulatory cycle driven by CARMIL and V-1 may promote actin network assembly at protruding edges. Proc. Natl. Acad. Sci. USA. 111:E1970–E1979.

    Not in our library10.1073/pnas.1313738111

  61. 61.

    Funk, J., F. Merino, M. Schaks, K. Rottner, S. Raunser, and P. Bieling. 2021. A barbed end interference mechanism reveals how capping protein promotes nucleation in branched actin networks. Nat. Commun. 12:5329.

    Not in our library10.1038/s41467-021-25682-5

  62. 62.

    Galkin, V.E., A. Orlova, W. Brieher, H.Y. Kueh, T.J. Mitchison, and E.H. Egelman. 2008. Coronin-1A stabilizes F-actin by bridging adjacent actin protomers and stapling opposite strands of the actin filament. J. Mol. Biol. 376:607–613.

    Not in our library10.1016/j.jmb.2007.12.007

  63. 63.

    Galkin, V.E., A. Orlova, D.S. Kudryashov, A. Solodukhin, E. Reisler, G.F. Schröder, and E.H. Egelman. 2011. Remodeling of actin filaments by ADF/cofilin proteins. Proc. Natl. Acad. Sci. USA. 108:20568–20572.

    Not in our library10.1073/pnas.1110109108

  64. 64.

    Galkin, V.E., A. Orlova, G.F. Schröder, and E.H. Egelman. 2010. Structural polymorphism in F-actin. Nat. Struct. Mol. Biol. 17:1318–1323.

    Not in our library10.1038/nsmb.1930

  65. 65.

    Gandhi, M., V. Achard, L. Blanchoin, and B.L. Goode. 2009. Coronin switches roles in actin disassembly depending on the nucleotide state of actin. Mol. Cell. 34:364–374.

    Not in our library10.1016/j.molcel.2009.02.029

  66. 66.

    Gandhi, M., and B.L. Goode. 2008. Coronin: The double-edged sword of actin dynamics. Subcell. Biochem. 48:72–87.

    Not in our library10.1007/978-0-387-09595-0_7

  67. 67.

    Gandhi, M., B.A. Smith, M. Bovellan, V. Paavilainen, K. Daugherty-Clarke, J. Gelles, P. Lappalainen, and B.L. Goode. 2010. GMF is a cofilin homolog that binds Arp2/3 complex to stimulate filament debranching and inhibit actin nucleation. Curr. Biol. 20:861–867.

    Not in our library10.1016/j.cub.2010.03.026

  68. 68.

    Gateva, G., E. Kremneva, T. Reindl, T. Kotila, K. Kogan, L. Gressin, P.W. Gunning, D.J. Manstein, A. Michelot, and P. Lappalainen. 2017. Tropomyosin isoforms specify functionally distinct actin filament populations in vitro. Curr. Biol. 27:705–713.

    Not in our library10.1016/j.cub.2017.01.018

  69. 69.

    Gatfield, J., I. Albrecht, B. Zanolari, M.O. Steinmetz, and J. Pieters. 2005. Association of the leukocyte plasma membrane with the actin cytoskeleton through coiled coil-mediated trimeric coronin 1 molecules. Mol. Biol. Cell. 16:2786–2798.

    Not in our library10.1091/mbc.e05-01-0042

  70. 70.

    Ge, P., Z.A.O. Durer, D. Kudryashov, Z.H. Zhou, and E. Reisler. 2014. Cryo-EM reveals different coronin binding modes for ADP- and ADP-BeFx actin filaments. Nat. Struct. Mol. Biol. 21:1075–1081.

    Not in our library10.1038/nsmb.2907

  71. 71.

    Ghosh, M., X. Song, G. Mouneimne, M. Sidani, D.S. Lawrence, and J.S. Condeelis. 2004. Cofilin promotes actin polymerization and defines the direction of cell motility. Science. 304:743–746.

    Not in our library10.1126/science.1094561

  72. 72.

    Giridharan, S.S.P., J.L. Rohn, N. Naslavsky, and S. Caplan. 2012. Differential regulation of actin microfilaments by human MICAL proteins. J. Cell Sci. 125:614–624.

    Not in our library10.1242/jcs.089367

  73. 73.

    Gonzalez Rodriguez, S., A.C.E. Wirshing, A.L. Goodman, and B.L. Goode. 2023. Cytosolic concentrations of actin binding proteins and the implications for in vivo F-actin turnover. J. Cell Biol. 222:e202306036.

    Not in our library10.1083/jcb.202306036

  74. 74.

    Goode, B.L., D.G. Drubin, and P. Lappalainen. 1998. Regulation of the cortical actin cytoskeleton in budding yeast by twinfilin, a ubiquitous actin monomer-sequestering protein. J. Cell Biol. 142:723–733.

    Not in our library10.1083/jcb.142.3.723

  75. 75.

    Goode, B.L., M.O. Sweeney, and J.A. Eskin. 2018. GMF as an actin network remodeling factor. Trends Cell Biol. 28:749–760.

    Not in our library10.1016/j.tcb.2018.04.008

  76. 76.

    Goode, B.L., J.J. Wong, A.C. Butty, M. Peter, A.L. McCormack, J.R. Yates, D.G. Drubin, and G. Barnes. 1999. Coronin promotes the rapid assembly and cross-linking of actin filaments and may link the actin and microtubule cytoskeletons in yeast. J. Cell Biol. 144:83–98.

    Not in our library10.1083/jcb.144.1.83

  77. 77.

    Goroncy, A.K., S. Koshiba, N. Tochio, T. Tomizawa, M. Sato, M. Inoue, S. Watanabe, Y. Hayashizaki, A. Tanaka, T. Kigawa, and S. Yokoyama. 2009. NMR solution structures of actin depolymerizing factor homology domains. Protein Sci. 18:2384–2392.

    Not in our library10.1002/pro.248

  78. 78.

    Gressin, L., A. Guillotin, C. Guérin, L. Blanchoin, and A. Michelot. 2015. Architecture dependence of actin filament network disassembly. Curr. Biol. 25:1437–1447.

    Not in our library10.1016/j.cub.2015.04.011

  79. 79.

    Grigoriev, I., K.L. Yu, E. Martinez-Sanchez, A. Serra-Marques, I. Smal, E. Meijering, J. Demmers, J. Peränen, R.J. Pasterkamp, P. van der Sluijs,. 2011. Rab6, Rab8, and MICAL3 cooperate in controlling docking and fusion of exocytotic carriers. Curr. Biol. 21:967–974.

    Not in our library10.1016/j.cub.2011.04.030

  80. 80.

    Grintsevich, E.E., G. Ahmed, A.A. Ginosyan, H. Wu, S.K. Rich, E. Reisler, and J.R. Terman. 2021. Profilin and Mical combine to impair F-actin assembly and promote disassembly and remodeling. Nat. Commun. 12:5542.

    Not in our library10.1038/s41467-021-25781-3

  81. 81.

    Grintsevich, E.E., P. Ge, M.R. Sawaya, H.G. Yesilyurt, J.R. Terman, Z.H. Zhou, and E. Reisler. 2017. Catastrophic disassembly of actin filaments via Mical-mediated oxidation. Nat. Commun. 8:2183.

    Not in our library10.1038/s41467-017-02357-8

  82. 82.

    Grintsevich, E.E., and E. Reisler. 2014. Drebrin inhibits cofilin-induced severing of F-actin. Cytoskeleton. 71:472–483.

    Not in our library10.1002/cm.21184

  83. 83.

    Grintsevich, E.E., H.G. Yesilyurt, S.K. Rich, R.J. Hung, J.R. Terman, and E. Reisler. 2016. F-actin dismantling through a redox-driven synergy between Mical and cofilin. Nat. Cell Biol. 18:876–885.

    Not in our library10.1038/ncb3390

  84. 84.

    Gunning, P.W., E.C. Hardeman, P. Lappalainen, and D.P. Mulvihill. 2015. Tropomyosin: Master regulator of actin filament function in the cytoskeleton. J. Cell Sci. 128:2965–2974.

    Not in our library10.1242/jcs.172502

  85. 85.

    Gunning, P.W., G. Schevzov, A.J. Kee, and E.C. Hardeman. 2005. Tropomyosin isoforms: Divining rods for actin cytoskeleton function. Trends Cell Biol. 15:333–341.

    Not in our library10.1016/j.tcb.2005.04.007

  86. 86.

    Guo, S., G.J. Hoeprich, J.O. Magliozzi, J. Gelles, and B.L. Goode. 2023. Dynamic remodeling of actin networks by cyclase-associated protein and CAP-Abp1 complexes. Curr. Biol. 33:4484–4495.e5.

    Not in our library10.1016/j.cub.2023.09.032

  87. 87.

    Guo, S., O.S. Sokolova, J. Chung, S. Padrick, J. Gelles, and B.L. Goode. 2018. Abp1 promotes Arp2/3 complex-dependent actin nucleation and stabilizes branch junctions by antagonizing GMF. Nat. Commun. 9:2895.

    Not in our library10.1038/s41467-018-05260-y

  88. 88.

    Hakala, M., H. Wioland, M. Tolonen, T. Kotila, A. Jegou, G. Romet-Lemonne, and P. Lappalainen. 2021. Twinfilin uncaps filament barbed ends to promote turnover of lamellipodial actin networks. Nat. Cell Biol. 23:147–159.

    Not in our library10.1038/s41556-020-00629-y

  89. 89.

    Hannappel, E., and W. Leibold. 1985. Biosynthesis rates and content of thymosin beta 4 in cell lines. Arch. Biochem. Biophys. 240:236–241.

    Not in our library10.1016/0003-9861(85)90028-1

  90. 90.

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

    Not in our library10.1074/jbc.m312718200

  91. 91.

    Haynes, E.M., S.B. Asokan, S.J. King, H.E. Johnson, J.M. Haugh, and J.E. Bear. 2015. GMFβ controls branched actin content and lamellipodial retraction in fibroblasts. J. Cell Biol. 209:803–812.

    Not in our library10.1083/jcb.201501094

  92. 92.

    Heimsath, E.G. Jr., and H.N. Higgs. 2012. The C terminus of formin FMNL3 accelerates actin polymerization and contains a WH2 domain-like sequence that binds both monomers and filament barbed ends. J. Biol. Chem. 287:3087–3098.

    Not in our library10.1074/jbc.m111.312207

  93. 93.

    Helfer, E., E.M. Nevalainen, P. Naumanen, S. Romero, D. Didry, D. Pantaloni, P. Lappalainen, and M.F. Carlier. 2006. Mammalian twinfilin sequesters ADP-G-actin and caps filament barbed ends: Implications in motility. EMBO J. 25:1184–1195.

    Not in our library10.1038/sj.emboj.7601019

  94. 94.

    Helgeson, L.A., and B.J. Nolen. 2013. Mechanism of synergistic activation of Arp2/3 complex by cortactin and N-WASP. Elife. 2:e00884.

    Not in our library10.7554/elife.00884

  95. 95.

    Hernandez-Valladares, M., T. Kim, B. Kannan, A. Tung, A.H. Aguda, M. Larsson, J.A. Cooper, and R.C. Robinson. 2010. Structural characterization of a capping protein interaction motif defines a family of actin filament regulators. Nat. Struct. Mol. Biol. 17:497–503.

    Not in our library10.1038/nsmb.1792

  96. 96.

    Hertzog, M., F. Milanesi, L. Hazelwood, A. Disanza, H. Liu, E. Perlade, M.G. Malabarba, S. Pasqualato, A. Maiolica, S. Confalonieri,. 2010. Molecular basis for the dual function of Eps8 on actin dynamics: Bundling and capping. PLoS Biol. 8:e1000387.

    Not in our library10.1371/journal.pbio.1000387

  97. 97.

    Hild, G., B. Bugyi, and M. Nyitrai. 2010. Conformational dynamics of actin: Effectors and implications for biological function. Cytoskeleton. 67:609–629.

    Not in our library10.1002/cm.20473

  98. 98.

    Hilton, D.M., R.M. Aguilar, A.B. Johnston, and B.L. Goode. 2018. Species-specific functions of twinfilin in actin filament depolymerization. J. Mol. Biol. 430:3323–3336.

    Not in our library10.1016/j.jmb.2018.06.025

  99. 99.

    Hliscs, M., J.M. Sattler, W. Tempel, J.D. Artz, A. Dong, R. Hui, K. Matuschewski, and H. Schüler. 2010. Structure and function of a G-actin sequestering protein with a vital role in malaria oocyst development inside the mosquito vector. J. Biol. Chem. 285:11572–11583.

    Not in our library10.1074/jbc.m109.054916

  100. 100.

    Hochmann, H., S. Pust, G. von Figura, K. Aktories, and H. Barth. 2006. Salmonella enterica SpvB ADP-ribosylates actin at position arginine-177-characterization of the catalytic domain within the SpvB protein and a comparison to binary clostridial actin-ADP-ribosylating toxins. Biochemistry. 45:1271–1277.

    Not in our library10.1021/bi051810w

  101. 101.

    Hoeprich, G.J., A.N. Sinclair, S. Shekhar, and B.L. Goode. 2022. Single-molecule imaging of IQGAP1 regulating actin filament dynamics. Mol. Biol. Cell. 33:ar2.

    Not in our library10.1091/mbc.e21-04-0211

  102. 102.

    Hotulainen, P., E. Paunola, M.K. Vartiainen, and P. Lappalainen. 2005. Actin-depolymerizing factor and cofilin-1 play overlapping roles in promoting rapid F-actin depolymerization in mammalian nonmuscle cells. Mol. Biol. Cell. 16:649–664.

    Not in our library10.1091/mbc.e04-07-0555

  103. 103.

    Huehn, A., W. Cao, W.A. Elam, X. Liu, E.M. De La Cruz, and C.V. Sindelar. 2018. The actin filament twist changes abruptly at boundaries between bare and cofilin-decorated segments. J. Biol. Chem. 293:5377–5383.

    Not in our library10.1074/jbc.ac118.001843

  104. 104.

    Humphries, C.L., H.I. Balcer, J.L. D’Agostino, B. Winsor, D.G. Drubin, G. Barnes, B.J. Andrews, and B.L. Goode. 2002. Direct regulation of Arp2/3 complex activity and function by the actin binding protein coronin. J. Cell Biol. 159:993–1004.

    Not in our library10.1083/jcb.200206113

  105. 105.

    Hung, R.J., C.W. Pak, and J.R. Terman. 2011. Direct redox regulation of F-actin assembly and disassembly by Mical. Science. 334:1710–1713.

    Not in our library10.1126/science.1211956

  106. 106.

    Hung, R.J., C.S. Spaeth, H.G. Yesilyurt, and J.R. Terman. 2013. SelR reverses Mical-mediated oxidation of actin to regulate F-actin dynamics. Nat. Cell Biol. 15:1445–1454.

    Not in our library10.1038/ncb2871

  107. 107.

    Hung, R.J., U. Yazdani, J. Yoon, H. Wu, T. Yang, N. Gupta, Z. Huang, W.J.H. van Berkel, and J.R. Terman. 2010. Mical links semaphorins to F-actin disassembly. Nature. 463:823–827.

    Not in our library10.1038/nature08724

  108. 108.

    Husson, C., L. Renault, D. Didry, D. Pantaloni, and M.F. Carlier. 2011. Cordon-Bleu uses WH2 domains as multifunctional dynamizers of actin filament assembly. Mol. Cell. 43:464–477.

    Not in our library10.1016/j.molcel.2011.07.010

  109. 109.

    Ikeda, K., R.K. Kundu, S. Ikeda, M. Kobara, H. Matsubara, and T. Quertermous. 2006. Glia maturation factor-gamma is preferentially expressed in microvascular endothelial and inflammatory cells and modulates actin cytoskeleton reorganization. Circ. Res. 99:424–433.

    Not in our library10.1161/01.res.0000237662.23539.0b

  110. 110.

    Inada, N. 2017. Plant actin depolymerizing factor: Actin microfilament disassembly and more. J. Plant Res. 130:227–238.

    Not in our library10.1007/s10265-016-0899-8

  111. 111.

    Ingerman, E., J.Y. Hsiao, and R.D. Mullins. 2013. Arp2/3 complex ATP hydrolysis promotes lamellipodial actin network disassembly but is dispensable for assembly. J. Cell Biol. 200:619–633.

    Not in our library10.1083/jcb.201211069

  112. 112.

    Ishikawa-Ankerhold, H.C., G. Gerisch, and A. Müller-Taubenberger. 2010. Genetic evidence for concerted control of actin dynamics in cytokinesis, endocytic traffic, and cell motility by coronin and Aip1. Cytoskeleton. 67:442–455.

    Not in our library10.1002/cm.20456

  113. 113.

    Iwasa, J.H., and R.D. Mullins. 2007. Spatial and temporal relationships between actin-filament nucleation, capping, and disassembly. Curr. Biol. 17:395–406.

    Not in our library10.1016/j.cub.2007.02.012

  114. 114.

    Jansen, S., A. Collins, S.M. Chin, C.A. Ydenberg, J. Gelles, and B.L. Goode. 2015. Single-molecule imaging of a three-component ordered actin disassembly mechanism. Nat. Commun. 6:7202.

    Not in our library10.1038/ncomms8202

  115. 115.

    Jansen, S., A. Collins, L. Golden, O. Sokolova, and B.L. Goode. 2014. Structure and mechanism of mouse cyclase-associated protein (CAP1) in regulating actin dynamics. J. Biol. Chem. 289:30732–30742.

    Not in our library10.1074/jbc.m114.601765

  116. 116.

    Jansen, S., and B.L. Goode. 2019. Tropomyosin isoforms differentially tune actin filament length and disassembly. Mol. Biol. Cell. 30:671–679.

    Not in our library10.1091/mbc.e18-12-0815

  117. 117.
    Individual Actin Filaments in a Microfluidic Flow Reveal the Mechanism of ATP Hydrolysis and Give Insight Into the Properties of Profilin · 2011
    No notice
  118. 118.

    Johnston, A.B., A. Collins, and B.L. Goode. 2015. High-speed depolymerization at actin filament ends jointly catalysed by Twinfilin and Srv2/CAP. Nat. Cell Biol. 17:1504–1511.

    Not in our library10.1038/ncb3252

  119. 119.

    Johnston, A.B., D.M. Hilton, P. McConnell, B. Johnson, M.T. Harris, A. Simone, G.K. Amarasinghe, J.A. Cooper, and B.L. Goode. 2018. A novel mode of capping protein-regulation by twinfilin. Elife. 7:e41313.

    Not in our library10.7554/elife.41313

  120. 120.

    Kaksonen, M., C.P. Toret, and D.G. Drubin. 2005. A modular design for the clathrin- and actin-mediated endocytosis machinery. Cell. 123:305–320.

    Not in our library10.1016/j.cell.2005.09.024

  121. 121.

    Kallio, J.P., and I. Kursula. 2014. Recombinant production, purification and crystallization of the Toxoplasma gondii coronin WD40 domain. Acta Crystallogr. F Struct. Biol. Commun. 70:517–521.

    Not in our library10.1107/s2053230x14005196

  122. 122.

    Kang, H., M.J. Bradley, W. Cao, K. Zhou, E.E. Grintsevich, A. Michelot, C.V. Sindelar, M. Hochstrasser, and E.M. De La Cruz. 2014. Site-specific cation release drives actin filament severing by vertebrate cofilin. Proc. Natl. Acad. Sci. USA. 111:17821–17826.

    Not in our library10.1073/pnas.1413397111

  123. 123.

    Karakozova, M., M. Kozak, C.C.L. Wong, A.O. Bailey, J.R. Yates III, A. Mogilner, H. Zebroski, and A. Kashina. 2006. Arginylation of beta-actin regulates actin cytoskeleton and cell motility. Science. 313:192–196.

    Not in our library10.1126/science.1129344

  124. 124.

    Karlsson, R., and P. Dráber. 2021. Profilin-A master coordinator of actin and microtubule organization in mammalian cells. J. Cell. Physiol. 236:7256–7265.

    Not in our library10.1002/jcp.30379

  125. 125.

    Kim, J., M.J. Shapiro, A.O. Bamidele, P. Gurel, P. Thapa, H.N. Higgs, K.E. Hedin, V.S. Shapiro, and D.D. Billadeau. 2014. Coactosin-like 1 antagonizes cofilin to promote lamellipodial protrusion at the immune synapse. PLoS One. 9:e85090.

    Not in our library10.1371/journal.pone.0085090

  126. 126.

    Kim, K., A. Yamashita, M.A. Wear, Y. Maéda, and J.A. Cooper. 2004. Capping protein binding to actin in yeast: Biochemical mechanism and physiological relevance. J. Cell Biol. 164:567–580.

    Not in our library10.1083/jcb.200308061

  127. 127.

    Kodera, N., H. Abe, P.D.N. Nguyen, and S. Ono. 2021. Native cyclase-associated protein and actin from Xenopus laevis oocytes form a unique 4:4 complex with a tripartite structure. J. Biol. Chem. 296:100649.

    Not in our library10.1016/j.jbc.2021.100649

  128. 128.
    F- and G-Actin Concentrations in Lamellipodia of Moving Cells · 2009
    No notice
  129. 129.

    Kotila, T., K. Kogan, G. Enkavi, S. Guo, I. Vattulainen, B.L. Goode, and P. Lappalainen. 2018. Structural basis of actin monomer re-charging by cyclase-associated protein. Nat. Commun. 9:1892.

    Not in our library10.1038/s41467-018-04231-7

  130. 130.
    Mechanism of synergistic actin filament pointed end depolymerization by cyclase-associated protein and cofilin · 2019
    No notice
  131. 131.
    Structural basis of rapid actin dynamics in the evolutionarily divergent Leishmania parasite · 2022
    Not checked
  132. 132.

    Kremneva, E., M.H. Makkonen, A. Skwarek-Maruszewska, G. Gateva, A. Michelot, R. Dominguez, and P. Lappalainen. 2014. Cofilin-2 controls actin filament length in muscle sarcomeres. Dev. Cell. 31:215–226.

    Not in our library10.1016/j.devcel.2014.09.002

  133. 133.

    Ksiazek, D., H. Brandstetter, L. Israel, G.P. Bourenkov, G. Katchalova, K.P. Janssen, H.D. Bartunik, A.A. Noegel, M. Schleicher, and T.A. Holak. 2003. Structure of the N-terminal domain of the adenylyl cyclase-associated protein (CAP) from Dictyostelium discoideum. Structure. 11:1171–1178.

    Not in our library10.1016/s0969-2126(03)00180-1

  134. 134.

    Kudryashova, E., H. Ankita, H. Ulrichs, S. Shekhar, and D.S. Kudryashov. 2022. Pointed-end processive elongation of actin filaments by Vibrio effectors VopF and VopL. Sci. Adv. 8:eadc9239.

    Not in our library10.1126/sciadv.adc9239

  135. 135.
    Glutamyl phosphate is an activated intermediate in actin crosslinking by actin crosslinking domain (ACD) toxin. · 2012
    Not checked
  136. 136.

    Kueh, H.Y., G.T. Charras, T.J. Mitchison, and W.M. Brieher. 2008. Actin disassembly by cofilin, coronin, and Aip1 occurs in bursts and is inhibited by barbed-end cappers. J. Cell Biol. 182:341–353.

    Not in our library10.1083/jcb.200801027

  137. 137.

    Kuhlman, P.A., C.A. Hughes, V. Bennett, and V.M. Fowler. 1996. A new function for adducin. Calcium/calmodulin-regulated capping of the barbed ends of actin filaments. J. Biol. Chem. 271:7986–7991.

    Not in our library10.1074/jbc.271.14.7986

  138. 138.

    Lacy, M.M., D. Baddeley, and J. Berro. 2019. Single-molecule turnover dynamics of actin and membrane coat proteins in clathrin-mediated endocytosis. Elife. 8:e52355.

    Not in our library10.7554/elife.52355

  139. 139.

    Lappalainen, P., and D.G. Drubin. 1997. Cofilin promotes rapid actin filament turnover in vivo. Nature. 388:78–82.

    Not in our library10.1038/40418

  140. 140.

    Lappalainen, P., E.V. Fedorov, A.A. Fedorov, S.C. Almo, and D.G. Drubin. 1997. Essential functions and actin-binding surfaces of yeast cofilin revealed by systematic mutagenesis. EMBO J. 16:5520–5530.

    Not in our library10.1093/emboj/16.18.5520

  141. 141.

    Lappalainen, P., M.M. Kessels, M.J. Cope, and D.G. Drubin. 1998. The ADF homology (ADF-H) domain: A highly exploited actin-binding module. Mol. Biol. Cell. 9:1951–1959.

    Not in our library10.1091/mbc.9.8.1951

  142. 142.

    Liang, Y., H. Niederstrasser, M. Edwards, C.E. Jackson, and J.A. Cooper. 2009. Distinct roles for CARMIL isoforms in cell migration. Mol. Biol. Cell. 20:5290–5305.

    Not in our library10.1091/mbc.e08-10-1071

  143. 143.

    Lim, R., D.E. Turriff, S.S. Troy, B.W. Moore, and L.F. Eng. 1977. Glia maturation factor: Effect on chemical differentiation of glioblasts in culture. Science. 195:195–196.

    Not in our library10.1126/science.188136

  144. 144.

    Lin, M.C., B.J. Galletta, D. Sept, and J.A. Cooper. 2010. Overlapping and distinct functions for cofilin, coronin and Aip1 in actin dynamics in vivo. J. Cell Sci. 123:1329–1342.

    Not in our library10.1242/jcs.065698

  145. 145.

    Loisel, T.P., R. Boujemaa, D. Pantaloni, and M.F. Carlier. 1999. Reconstitution of actin-based motility of Listeria and Shigella using pure proteins. Nature. 401:613–616.

    Not in our library10.1038/44183

  146. 146.

    Luan, Q., and B.J. Nolen. 2013. Structural basis for regulation of Arp2/3 complex by GMF. Nat. Struct. Mol. Biol. 20:1062–1068.

    Not in our library10.1038/nsmb.2628

  147. 147.

    Mabuchi, I. 1983. An actin-depolymerizing protein (depactin) from starfish oocytes: Properties and interaction with actin. J. Cell Biol. 97:1612–1621.

    Not in our library10.1083/jcb.97.5.1612

  148. 148.

    Makkonen, M., E. Bertling, N.A. Chebotareva, J. Baum, and P. Lappalainen. 2013. Mammalian and malaria parasite cyclase-associated proteins catalyze nucleotide exchange on G-actin through a conserved mechanism. J. Biol. Chem. 288:984–994.

    Not in our library10.1074/jbc.m112.435719

  149. 149.

    Mallavarapu, A., and T. Mitchison. 1999. Regulated actin cytoskeleton assembly at filopodium tips controls their extension and retraction. J. Cell Biol. 146:1097–1106.

    Not in our library10.1083/jcb.146.5.1097

  150. 150.

    Martin, A.C., M.D. Welch, and D.G. Drubin. 2006. Arp2/3 ATP hydrolysis-catalysed branch dissociation is critical for endocytic force generation. Nat. Cell Biol. 8:826–833.

    Not in our library10.1038/ncb1443

  151. 151.

    Martin, A.C., X.P. Xu, I. Rouiller, M. Kaksonen, Y. Sun, L. Belmont, N. Volkmann, D. Hanein, M. Welch, and D.G. Drubin. 2005. Effects of Arp2 and Arp3 nucleotide-binding pocket mutations on Arp2/3 complex function. J. Cell Biol. 168:315–328.

    Not in our library10.1083/jcb.200408177

  152. 152.

    Mattila, P.K., O. Quintero-Monzon, J. Kugler, J.B. Moseley, S.C. Almo, P. Lappalainen, and B.L. Goode. 2004. A high-affinity interaction with ADP-actin monomers underlies the mechanism and in vivo function of Srv2/cyclase-associated protein. Mol. Biol. Cell. 15:5158–5171.

    Not in our library10.1091/mbc.e04-06-0444

  153. 153.

    Mehta, S., and L.D. Sibley. 2010. Toxoplasma gondii actin depolymerizing factor acts primarily to sequester G-actin. J. Biol. Chem. 285:6835–6847.

    Not in our library10.1074/jbc.m109.068155

  154. 154.

    Mejillano, M.R., S.i. Kojima, D.A. Applewhite, F.B. Gertler, T.M. Svitkina, and G.G. Borisy. 2004. Lamellipodial versus filopodial mode of the actin nanomachinery: Pivotal role of the filament barbed end. Cell. 118:363–373.

    Not in our library10.1016/j.cell.2004.07.019

  155. 155.

    Mellor, H. 2010. The role of formins in filopodia formation. Biochim. Biophys. Acta. 1803:191–200.

    Not in our library10.1016/j.bbamcr.2008.12.018

  156. 156.

    Mikati, M.A., D. Breitsprecher, S. Jansen, E. Reisler, and B.L. Goode. 2015. Coronin enhances actin filament severing by recruiting cofilin to filament sides and altering F-actin conformation. J. Mol. Biol. 427:3137–3147.

    Not in our library10.1016/j.jmb.2015.08.011

  157. 157.

    Miyoshi, T., T. Tsuji, C. Higashida, M. Hertzog, A. Fujita, S. Narumiya, G. Scita, and N. Watanabe. 2006. Actin turnover-dependent fast dissociation of capping protein in the dendritic nucleation actin network: Evidence of frequent filament severing. J. Cell Biol. 175:947–955.

    Not in our library10.1083/jcb.200604176

  158. 158.

    Mockrin, S.C., and E.D. Korn. 1980. Acanthamoeba profilin interacts with G-actin to increase the rate of exchange of actin-bound adenosine 5′-triphosphate. Biochemistry. 19:5359–5362.

    Not in our library10.1021/bi00564a033

  159. 159.

    Mohri, K., S. Vorobiev, A.A. Fedorov, S.C. Almo, and S. Ono. 2004. Identification of functional residues on Caenorhabditis elegans actin-interacting protein 1 (UNC-78) for disassembly of actin depolymerizing factor/cofilin-bound actin filaments. J. Biol. Chem. 279:31697–31707.

    Not in our library10.1074/jbc.m403351200

  160. 160.

    Moriyama, K., and I. Yahara. 2002. Human CAP1 is a key factor in the recycling of cofilin and actin for rapid actin turnover. J. Cell Sci. 115:1591–1601.

    Not in our library10.1242/jcs.115.8.1591

  161. 161.

    Mullins, R.D., J.A. Heuser, and T.D. Pollard. 1998. 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. 95:6181–6186.

    Not in our library10.1073/pnas.95.11.6181

  162. 162.

    Mwangangi, D.M., E. Manser, and R.C. Robinson. 2021. The structure of the actin filament uncapping complex mediated by twinfilin. Sci. Adv. 7:eabd5271.

    Not in our library10.1126/sciadv.abd5271

  163. 163.

    Nadkarni, A.V., and W.M. Brieher. 2014. Aip1 destabilizes cofilin-saturated actin filaments by severing and accelerating monomer dissociation from ends. Curr. Biol. 24:2749–2757.

    Not in our library10.1016/j.cub.2014.09.048

  164. 164.

    Nag, S., M. Larsson, R.C. Robinson, and L.D. Burtnick. 2013. Gelsolin: The tail of a molecular gymnast. Cytoskeleton. 70:360–384.

    Not in our library10.1002/cm.21117

  165. 165.

    Nakano, K., H. Kuwayama, M. Kawasaki, O. Numata, and M. Takaine. 2010. GMF is an evolutionarily developed Adf/cofilin-super family protein involved in the Arp2/3 complex-mediated organization of the actin cytoskeleton. Cytoskeleton. 67:373–382.

    Not in our library10.1002/cm.20451

  166. 166.

    Nakano, K., and I. Mabuchi. 2006. Actin-depolymerizing protein Adf1 is required for formation and maintenance of the contractile ring during cytokinesis in fission yeast. Mol. Biol. Cell. 17:1933–1945.

    Not in our library10.1091/mbc.e05-09-0900

  167. 167.

    Narayanan, P., P. Chatterton, A. Ikeda, S. Ikeda, D.P. Corey, J.M. Ervasti, and B.J. Perrin. 2015. Length regulation of mechanosensitive stereocilia depends on very slow actin dynamics and filament-severing proteins. Nat. Commun. 6:6855.

    Not in our library10.1038/ncomms7855

  168. 168.
    Cofilin-induced unidirectional cooperative conformational changes in actin filaments revealed by high-speed atomic force microscopy · 2015
    No notice
  169. 169.

    Nishida, E. 1985. Opposite effects of cofilin and profilin from porcine brain on rate of exchange of actin-bound adenosine 5′-triphosphate. Biochemistry. 24:1160–1164.

    Not in our library10.1021/bi00326a015

  170. 170.

    Nomura, K., and S. Ono. 2013. ATP-dependent regulation of actin monomer-filament equilibrium by cyclase-associated protein and ADF/cofilin. Biochem. J. 453:249–259.

    Not in our library10.1042/bj20130491

  171. 171.

    Normoyle, K.P.M., and W.M. Brieher. 2012. Cyclase-associated protein (CAP) acts directly on F-actin to accelerate cofilin-mediated actin severing across the range of physiological pH. J. Biol. Chem. 287:35722–35732.

    Not in our library10.1074/jbc.m112.396051

  172. 172.

    Ojala, P.J., V.O. Paavilainen, M.K. Vartiainen, R. Tuma, A.G. Weeds, and P. Lappalainen. 2002. The two ADF-H domains of twinfilin play functionally distinct roles in interactions with actin monomers. Mol. Biol. Cell. 13:3811–3821.

    Not in our library10.1091/mbc.e02-03-0157

  173. 173.

    Okabe, S., and N. Hirokawa. 1989. Incorporation and turnover of biotin-labeled actin microinjected into fibroblastic cells: An immunoelectron microscopic study. J. Cell Biol. 109:1581–1595.

    Not in our library10.1083/jcb.109.4.1581

  174. 174.

    Okada, K., L. Blanchoin, H. Abe, H. Chen, T.D. Pollard, and J.R. Bamburg. 2002. Xenopus actin-interacting protein 1 (XAip1) enhances cofilin fragmentation of filaments by capping filament ends. J. Biol. Chem. 277:43011–43016.

    Not in our library10.1074/jbc.m203111200

  175. 175.

    Okada, K., T. Obinata, and H. Abe. 1999. XAIP1: A Xenopus homologue of yeast actin interacting protein 1 (AIP1), which induces disassembly of actin filaments cooperatively with ADF/cofilin family proteins. J. Cell Sci. 112:1553–1565.

    Not in our library10.1242/jcs.112.10.1553

  176. 176.

    Okada, K., H. Ravi, E.M. Smith, and B.L. Goode. 2006. Aip1 and cofilin promote rapid turnover of yeast actin patches and cables: A coordinated mechanism for severing and capping filaments. Mol. Biol. Cell. 17:2855–2868.

    Not in our library10.1091/mbc.e06-02-0135

  177. 177.

    Okamoto, H., H. Fujita, S. Matsuyama, and S. Tsuyama. 1997. Purification, characterization, and localization of an ADP-ribosylactin hydrolase that uses ADP-ribosylated actin from rat brains as a substrate. J. Biol. Chem. 272:28116–28125.

    Not in our library10.1074/jbc.272.44.28116

  178. 178.

    Okreglak, V., and D.G. Drubin. 2007. Cofilin recruitment and function during actin-mediated endocytosis dictated by actin nucleotide state. J. Cell Biol. 178:1251–1264.

    Not in our library10.1083/jcb.200703092

  179. 179.

    Oku, T., S. Itoh, R. Ishii, K. Suzuki, W.M. Nauseef, S. Toyoshima, and T. Tsuji. 2005. Homotypic dimerization of the actin-binding protein p57/coronin-1 mediated by a leucine zipper motif in the C-terminal region. Biochem. J. 387:325–331.

    Not in our library10.1042/bj20041020

  180. 180.

    Ono, S. 2013. The role of cyclase-associated protein in regulating actin filament dynamics: More than a monomer-sequestration factor. J. Cell Sci. 126:3249–3258.

    Not in our library10.1242/jcs.128231

  181. 181.

    Ono, S. 2018. Functions of actin-interacting protein 1 (AIP1)/WD repeat protein 1 (WDR1) in actin filament dynamics and cytoskeletal regulation. Biochem. Biophys. Res. Commun. 506:315–322.

    Not in our library10.1016/j.bbrc.2017.10.096

  182. 182.

    Ostrowska, Z., K. Robaszkiewicz, and J. Moraczewska. 2017. Regulation of actin filament turnover by cofilin-1 and cytoplasmic tropomyosin isoforms. Biochim. Biophys. Acta Proteins Proteom. 1865:88–98.

    Not in our library10.1016/j.bbapap.2016.09.019

  183. 183.

    Paavilainen, V.O., M. Hellman, E. Helfer, M. Bovellan, A. Annila, M.F. Carlier, P. Permi, and P. Lappalainen. 2007. Structural basis and evolutionary origin of actin filament capping by twinfilin. Proc. Natl. Acad. Sci. USA. 104:3113–3118.

    Not in our library10.1073/pnas.0608725104

  184. 184.

    Palmgren, S., P.J. Ojala, M.A. Wear, J.A. Cooper, and P. Lappalainen. 2001. Interactions with PIP2, ADP-actin monomers, and capping protein regulate the activity and localization of yeast twinfilin. J. Cell Biol. 155:251–260.

    Not in our library10.1083/jcb.200106157

  185. 185.

    Pandit, N.G., W. Cao, J. Bibeau, E.M. Johnson-Chavarria, E.W. Taylor, T.D. Pollard, and E.M. De La Cruz. 2020. Force and phosphate release from Arp2/3 complex promote dissociation of actin filament branches. Proc. Natl. Acad. Sci. USA. 117:13519–13528.

    Not in our library10.1073/pnas.1911183117

  186. 186.

    Pantaloni, D., and M.F. Carlier. 1993. How profilin promotes actin filament assembly in the presence of thymosin beta 4. Cell. 75:1007–1014.

    Not in our library10.1016/0092-8674(93)90544-z

  187. 187.

    Pelikan-Conchaudron, A., C. Le Clainche, D. Didry, and M.F. Carlier. 2011. The IQGAP1 protein is a calmodulin-regulated barbed end capper of actin filaments: Possible implications in its function in cell migration. J. Biol. Chem. 286:35119–35128.

    Not in our library10.1074/jbc.m111.258772

  188. 188.

    Perelroizen, I., M.F. Carlier, and D. Pantaloni. 1995. Binding of divalent cation and nucleotide to G-actin in the presence of profilin. J. Biol. Chem. 270:1501–1508.

    Not in our library10.1074/jbc.270.4.1501

  189. 189.

    Perieteanu, A.A., D.D. Visschedyk, A.R. Merrill, and J.F. Dawson. 2010. ADP-ribosylation of cross-linked actin generates barbed-end polymerization-deficient F-actin oligomers. Biochemistry. 49:8944–8954.

    Not in our library10.1021/bi1008062

  190. 190.

    Pimm, M.L., A.G. Marcin, B.K. Haarer, M.A. Eligio, and J.L. Henty-Ridilla. 2023. Coordination of actin plus-end dynamics by IQGAP1, formin, and capping protein. bioRxiv. (Preprint posted May 04, 2023).

    Not in our library10.1101/2023.05.04.539490

  191. 191.

    Pollard, T.D. 1986. Rate constants for the reactions of ATP- and ADP-actin with the ends of actin filaments. J. Cell Biol. 103:2747–2754.

    Not in our library10.1083/jcb.103.6.2747

  192. 192.

    Pollard, T.D., and G.G. Borisy. 2003. Cellular motility driven by assembly and disassembly of actin filaments. Cell. 112:453–465.

    Not in our library10.1016/s0092-8674(03)00120-x

  193. 193.

    Pollard, T.D., and J.A. Cooper. 1984. Quantitative analysis of the effect of Acanthamoeba profilin on actin filament nucleation and elongation. Biochemistry. 23:6631–6641.

    Not in our library10.1021/bi00321a054

  194. 194.

    Pollard, T.D., and M.S. Mooseker. 1981. Direct measurement of actin polymerization rate constants by electron microscopy of actin filaments nucleated by isolated microvillus cores. J. Cell Biol. 88:654–659.

    Not in our library10.1083/jcb.88.3.654

  195. 195.

    Poukkula, M., M. Hakala, N. Pentinmikko, M.O. Sweeney, S. Jansen, J. Mattila, V. Hietakangas, B.L. Goode, and P. Lappalainen. 2014. GMF promotes leading-edge dynamics and collective cell migration in vivo. Curr. Biol. 24:2533–2540.

    Not in our library10.1016/j.cub.2014.08.066

  196. 196.

    Poukkula, M., E. Kremneva, M. Serlachius, and P. Lappalainen. 2011. Actin-depolymerizing factor homology domain: A conserved fold performing diverse roles in cytoskeletal dynamics. Cytoskeleton. 68:471–490.

    Not in our library10.1002/cm.20530

  197. 197.

    Purde, V., F. Busch, E. Kudryashova, V.H. Wysocki, and D.S. Kudryashov. 2019. Oligomerization affects the ability of human cyclase-associated proteins 1 and 2 to promote actin severing by cofilins. Int. J. Mol. Sci. 20:5647.

    Not in our library10.3390/ijms20225647

  198. 198.

    Quintero-Monzon, O., E.M. Jonasson, E. Bertling, L. Talarico, F. Chaudhry, M. Sihvo, P. Lappalainen, and B.L. Goode. 2009. Reconstitution and dissection of the 600-kD Srv2/CAP complex: Roles for oligomerization and cofilin-actin binding in driving actin turnover. J. Biol. Chem. 284:10923–10934.

    Not in our library10.1074/jbc.m808760200

  199. 199.

    Rao, J.N., Y. Madasu, and R. Dominguez. 2014. Mechanism of actin filament pointed-end capping by tropomodulin. Science. 345:463–467.

    Not in our library10.1126/science.1256159

  200. 200.

    Robaszkiewicz, K., Z. Ostrowska, K. Marchlewicz, and J. Moraczewska. 2016. Tropomyosin isoforms differentially modulate the regulation of actin filament polymerization and depolymerization by cofilins. FEBS J. 283:723–737.

    Not in our library10.1111/febs.13626

  201. 201.

    Rodal, A.A., L. Kozubowski, B.L. Goode, D.G. Drubin, and J.H. Hartwig. 2005. Actin and septin ultrastructures at the budding yeast cell cortex. Mol. Biol. Cell. 16:372–384.

    Not in our library10.1091/mbc.e04-08-0734

  202. 202.

    Rodal, A.A., J.W. Tetreault, P. Lappalainen, D.G. Drubin, and D.C. Amberg. 1999. Aip1p interacts with cofilin to disassemble actin filaments. J. Cell Biol. 145:1251–1264.

    Not in our library10.1083/jcb.145.6.1251

  203. 203.

    Rotty, J.D., C. Wu, and J.E. Bear. 2013. New insights into the regulation and cellular functions of the ARP2/3 complex. Nat. Rev. Mol. Cell Biol. 14:7–12.

    Not in our library10.1038/nrm3492

  204. 204.

    Rzadzinska, A.K., E.M. Nevalainen, H.M. Prosser, P. Lappalainen, and K.P. Steel. 2009. MyosinVIIa interacts with Twinfilin-2 at the tips of mechanosensory stereocilia in the inner ear. PLoS One. 4:e7097.

    Not in our library10.1371/journal.pone.0007097

  205. 205.

    Rzadzinska, A.K., M.E. Schneider, C. Davies, G.P. Riordan, and B. Kachar. 2004. An actin molecular treadmill and myosins maintain stereocilia functional architecture and self-renewal. J. Cell Biol. 164:887–897.

    Not in our library10.1083/jcb.200310055

  206. 206.

    Safer, D., M. Elzinga, and V.T. Nachmias. 1991. Thymosin beta 4 and Fx, an actin-sequestering peptide, are indistinguishable. J. Biol. Chem. 266:4029–4032.

    Not in our library10.1016/s0021-9258(20)64278-8

  207. 207.

    Sattler, J.M., M. Ganter, M. Hliscs, K. Matuschewski, and H. Schüler. 2011. Actin regulation in the malaria parasite. Eur. J. Cell Biol. 90:966–971.

    Not in our library10.1016/j.ejcb.2010.11.011

  208. 208.

    Shekhar, S., and M.F. Carlier. 2017. Enhanced depolymerization of actin filaments by ADF/cofilin and monomer funneling by capping protein cooperate to accelerate barbed-end growth. Curr. Biol. 27:1990–1998.e5.

    Not in our library10.1016/j.cub.2017.05.036

  209. 209.
    Synergy between Cyclase-associated protein and Cofilin accelerates actin filament depolymerization by two orders of magnitude · 2019
    Not checked
  210. 210.

    Shekhar, S., G.J. Hoeprich, J. Gelles, and B.L. Goode. 2021. Twinfilin bypasses assembly conditions and actin filament aging to drive barbed end depolymerization. J. Cell Biol. 220:e202006022.

    Not in our library10.1083/jcb.202006022

  211. 211.

    Shekhar, S., M. Kerleau, S. Kühn, J. Pernier, G. Romet-Lemonne, A. Jégou, and M.F. Carlier. 2015. Formin and capping protein together embrace the actin filament in a ménage à trois. Nat. Commun. 6:8730.

    Not in our library10.1038/ncomms9730

  212. 212.

    Shekhar, S., J. Pernier, and M.F. Carlier. 2016. Regulators of actin filament barbed ends at a glance. J. Cell Sci. 129:1085–1091.

    Not in our library10.1242/jcs.179994

  213. 213.

    Shi, M., Y. Xie, Y. Zheng, J. Wang, Y. Su, Q. Yang, and S. Huang. 2013. Oryza sativa actin-interacting protein 1 is required for rice growth by promoting actin turnover. Plant J. 73:747–760.

    Not in our library10.1111/tpj.12065

  214. 214.

    Sinnar, S.A., S. Antoku, J.M. Saffin, J.A. Cooper, and S. Halpain. 2014. Capping protein is essential for cell migration in vivo and for filopodial morphology and dynamics. Mol. Biol. Cell. 25:2152–2160.

    Not in our library10.1091/mbc.e13-12-0749

  215. 215.

    Smith, M.G., S.R. Swamy, and L.A. Pon. 2001. The life cycle of actin patches in mating yeast. J. Cell Sci. 114:1505–1513.

    Not in our library10.1242/jcs.114.8.1505

  216. 216.

    Sokolova, O.S., A. Chemeris, S. Guo, S.L. Alioto, M. Gandhi, S. Padrick, E. Pechnikova, V. David, A. Gautreau, and B.L. Goode. 2017. Structural basis of Arp2/3 complex inhibition by GMF, coronin, and arpin. J. Mol. Biol. 429:237–248.

    Not in our library10.1016/j.jmb.2016.11.030

  217. 217.

    Spoerl, Z., M. Stumpf, A.A. Noegel, and A. Hasse. 2002. Oligomerization, F-actin interaction, and membrane association of the ubiquitous mammalian coronin 3 are mediated by its carboxyl terminus. J. Biol. Chem. 277:48858–48867.

    Not in our library10.1074/jbc.m205136200

  218. 218.

    Suarez, C., J. Roland, R. Boujemaa-Paterski, H. Kang, B.R. McCullough, A.C. Reymann, C. Guérin, J.L. Martiel, E.M. De la Cruz, and L. Blanchoin. 2011. Cofilin tunes the nucleotide state of actin filaments and severs at bare and decorated segment boundaries. Curr. Biol. 21:862–868.

    Not in our library10.1016/j.cub.2011.03.064

  219. 219.

    Svitkina, T. 2018. The actin cytoskeleton and actin-based motility. Cold Spring Harb. Perspect. Biol. 10:a018267.

    Not in our library10.1101/cshperspect.a018267

  220. 220.

    Svitkina, T.M., and G.G. Borisy. 1999. Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J. Cell Biol. 145:1009–1026.

    Not in our library10.1083/jcb.145.5.1009

  221. 221.

    Svitkina, T.M., E.A. Bulanova, O.Y. Chaga, D.M. Vignjevic, S.i. Kojima, J.M. Vasiliev, and G.G. Borisy. 2003. Mechanism of filopodia initiation by reorganization of a dendritic network. J. Cell Biol. 160:409–421.

    Not in our library10.1083/jcb.200210174

  222. 222.

    Tang, V.W., A.V. Nadkarni, and W.M. Brieher. 2020. Catastrophic actin filament bursting by cofilin, Aip1, and coronin. J. Biol. Chem. 295:13299–13313.

    Not in our library10.1074/jbc.ra120.015018

  223. 223.

    Taoka, M., T. Ichimura, A. Wakamiya-Tsuruta, Y. Kubota, T. Araki, T. Obinata, and T. Isobe. 2003. V-1, a protein expressed transiently during murine cerebellar development, regulates actin polymerization via interaction with capping protein. J. Biol. Chem. 278:5864–5870.

    Not in our library10.1074/jbc.m211509200

  224. 224.

    Terman, J.R., and A. Kashina. 2013. Post-translational modification and regulation of actin. Curr. Opin. Cell Biol. 25:30–38.

    Not in our library10.1016/j.ceb.2012.10.009

  225. 225.

    Theriot, J.A., and T.J. Mitchison. 1991. Actin microfilament dynamics in locomoting cells. Nature. 352:126–131.

    Not in our library10.1038/352126a0

  226. 226.

    Theriot, J.A., and T.J. Mitchison. 1992. Comparison of actin and cell surface dynamics in motile fibroblasts. J. Cell Biol. 119:367–377.

    Not in our library10.1083/jcb.119.2.367

  227. 227.

    Theriot, J.A., T.J. Mitchison, L.G. Tilney, and D.A. Portnoy. 1992. The rate of actin-based motility of intracellular Listeria monocytogenes equals the rate of actin polymerization. Nature. 357:257–260.

    Not in our library10.1038/357257a0

  228. 228.

    Toshima, J.Y., C. Horikomi, A. Okada, M.N. Hatori, M. Nagano, A. Masuda, W. Yamamoto, D.E. Siekhaus, and J. Toshima. 2016. Srv2/CAP is required for polarized actin cable assembly and patch internalization during clathrin-mediated endocytosis. J. Cell Sci. 129:367–379.

    Not in our library10.1242/jcs.176651

  229. 229.

    Ulrichs, H., I. Gaska, and S. Shekhar. 2023. Multicomponent regulation of actin barbed end assembly by twinfilin, formin and capping protein. Nat. Commun. 14:3981.

    Not in our library10.1038/s41467-023-39655-3

  230. 230.

    Urban, E., S. Jacob, M. Nemethova, G.P. Resch, and J.V. Small. 2010. Electron tomography reveals unbranched networks of actin filaments in lamellipodia. Nat. Cell Biol. 12:429–435.

    Not in our library10.1038/ncb2044

  231. 231.

    Uruno, T., J. Liu, P. Zhang, C. Fan Yx, C. Egile, R. Li, S.C. Mueller, and X. Zhan. 2001. Activation of Arp2/3 complex-mediated actin polymerization by cortactin. Nat. Cell Biol. 3:259–266.

    Not in our library10.1038/35060051

  232. 232.

    Vandekerckhove, J., B. Schering, M. Bärmann, and K. Aktories. 1988. Botulinum C2 toxin ADP-ribosylates cytoplasmic beta/gamma-actin in arginine 177. J. Biol. Chem. 263:696–700.

    Not in our library10.1016/s0021-9258(19)35408-0

  233. 233.

    Vartiainen, M., P.J. Ojala, P. Auvinen, J. Peränen, and P. Lappalainen. 2000. Mouse A6/twinfilin is an actin monomer-binding protein that localizes to the regions of rapid actin dynamics. Mol. Cell. Biol. 20:1772–1783.

    Not in our library10.1128/mcb.20.5.1772-1783.2000

  234. 234.

    Vartiainen, M.K., T. Mustonen, P.K. Mattila, P.J. Ojala, I. Thesleff, J. Partanen, and P. Lappalainen. 2002. The three mouse actin-depolymerizing factor/cofilins evolved to fulfill cell-type-specific requirements for actin dynamics. Mol. Biol. Cell. 13:183–194.

    Not in our library10.1091/mbc.01-07-0331

  235. 235.

    Vartiainen, M.K., E.M. Sarkkinen, T. Matilainen, M. Salminen, and P. Lappalainen. 2003. Mammals have two twinfilin isoforms whose subcellular localizations and tissue distributions are differentially regulated. J. Biol. Chem. 278:34347–34355.

    Not in our library10.1074/jbc.m303642200

  236. 236.

    Vizcaíno-Castillo, A., J.F. Osorio-Méndez, J.R. Ambrosio, R. Hernández, and A.M. Cevallos. 2020. The complexity and diversity of the actin cytoskeleton of trypanosomatids. Mol. Biochem. Parasitol. 237:111278.

    Not in our library10.1016/j.molbiopara.2020.111278

  237. 237.

    Wang, J., J.M. Sanger, and J.W. Sanger. 2005. Differential effects of latrunculin-A on myofibrils in cultures of skeletal muscle cells: Insights into mechanisms of myofibrillogenesis. Cell Motil. Cytoskeleton. 62:35–47.

    Not in our library10.1002/cm.20083

  238. 238.

    Wang, Y.L. 1984. Reorganization of actin filament bundles in living fibroblasts. J. Cell Biol. 99:1478–1485.

    Not in our library10.1083/jcb.99.4.1478

  239. 239.

    Wang, Y.L. 1985. Exchange of actin subunits at the leading edge of living fibroblasts: Possible role of treadmilling. J. Cell Biol. 101:597–602.

    Not in our library10.1083/jcb.101.2.597

  240. 240.

    Watanabe, N., and T.J. Mitchison. 2002. Single-molecule speckle analysis of actin filament turnover in lamellipodia. Science. 295:1083–1086.

    Not in our library10.1126/science.1067470

  241. 241.

    Weaver, A.M., J.E. Heuser, A.V. Karginov, W.L. Lee, J.T. Parsons, and J.A. Cooper. 2002. Interaction of cortactin and N-WASp with Arp2/3 complex. Curr. Biol. 12:1270–1278.

    Not in our library10.1016/s0960-9822(02)01035-7

  242. 242.

    Weber, A., V.T. Nachmias, C.R. Pennise, M. Pring, and D. Safer. 1992. Interaction of thymosin beta 4 with muscle and platelet actin: Implications for actin sequestration in resting platelets. Biochemistry. 31:6179–6185.

    Not in our library10.1021/bi00142a002

  243. 243.

    Wegner, A., and K. Aktories. 1988. ADP-ribosylated actin caps the barbed ends of actin filaments. J. Biol. Chem. 263:13739–13742.

    Not in our library10.1016/s0021-9258(18)68303-6

  244. 244.

    Wegner, A., and G. Isenberg. 1983. 12-fold difference between the critical monomer concentrations of the two ends of actin filaments in physiological salt conditions. Proc. Natl. Acad. Sci. USA. 80:4922–4925.

    Not in our library10.1073/pnas.80.16.4922

  245. 245.

    Weigt, C., I. Just, A. Wegner, and K. Aktories. 1989. Nonmuscle actin ADP-ribosylated by botulinum C2 toxin caps actin filaments. FEBS Lett. 246:181–184.

    Not in our library10.1016/0014-5793(89)80279-0

  246. 246.

    Wioland, H., S. Frémont, B. Guichard, A. Echard, A. Jégou, and G. Romet-Lemonne. 2021. Actin filament oxidation by MICAL1 suppresses protections from cofilin-induced disassembly. EMBO Rep. 22:e50965.

    Not in our library10.15252/embr.202050965

  247. 247.

    Wioland, H., B. Guichard, Y. Senju, S. Myram, P. Lappalainen, A. Jegou, and G. Romet-Lemonne. 2017. ADF/cofilin accelerates actin dynamics by severing filaments and promoting their depolymerization at both ends. Curr. Biol. 27:1956–1967.e7.

    Not in our library10.1016/j.cub.2017.05.048

  248. 248.

    Wioland, H., A. Jegou, and G. Romet-Lemonne. 2019a. Quantitative variations with pH of actin depolymerizing factor/cofilin’s multiple actions on actin filaments. Biochemistry. 58:40–47.

    Not in our library10.1021/acs.biochem.8b01001

  249. 249.

    Wioland, H., A. Jegou, and G. Romet-Lemonne. 2019b. Torsional stress generated by ADF/cofilin on cross-linked actin filaments boosts their severing. Proc. Natl. Acad. Sci. USA. 116:2595–2602.

    Not in our library10.1073/pnas.1812053116

  250. 250.

    Wirshing, A.C.E., S.G. Rodriguez, and B.L. Goode. 2023. Evolutionary tuning of barbed end competition allows simultaneous construction of architecturally distinct actin structures. J. Cell Biol. 222:e202209105.

    Not in our library10.1083/jcb.202209105

  251. 251.

    Xie, C., Y. Jiang, Z. Zhu, S. Huang, W. Li, and G. Ou. 2021. Actin filament debranching regulates cell polarity during cell migration and asymmetric cell division. Proc. Natl. Acad. Sci. USA. 118:e2100805118.

    Not in our library10.1073/pnas.2100805118

  252. 252.

    Yang, C., M. Pring, M.A. Wear, M. Huang, J.A. Cooper, T.M. Svitkina, and S.H. Zigmond. 2005. Mammalian CARMIL inhibits actin filament capping by capping protein. Dev. Cell. 9:209–221.

    Not in our library10.1016/j.devcel.2005.06.008

  253. 253.

    Yang, C., and T. Svitkina. 2011. Filopodia initiation: Focus on the Arp2/3 complex and formins. Cell Adh. Migr. 5:402–408.

    Not in our library10.4161/cam.5.5.16971

  254. 254.

    Ydenberg, C.A., A. Johnston, J. Weinstein, D. Bellavance, S. Jansen, and B.L. Goode. 2015. Combinatorial genetic analysis of a network of actin disassembly-promoting factors. Cytoskeleton. 72:349–361.

    Not in our library10.1002/cm.21231

  255. 255.

    Ydenberg, C.A., S.B. Padrick, M.O. Sweeney, M. Gandhi, O. Sokolova, and B.L. Goode. 2013. GMF severs actin-Arp2/3 complex branch junctions by a cofilin-like mechanism. Curr. Biol. 23:1037–1045.

    Not in our library10.1016/j.cub.2013.04.058

  256. 256.

    Ydenberg, C.A., B.A. Smith, D. Breitsprecher, J. Gelles, and B.L. Goode. 2011. Cease-fire at the leading edge: New perspectives on actin filament branching, debranching, and cross-linking. Cytoskeleton. 68:596–602.

    Not in our library10.1002/cm.20543

  257. 257.

    Young, M.E., J.A. Cooper, and P.C. Bridgman. 2004. Yeast actin patches are networks of branched actin filaments. J. Cell Biol. 166:629–635.

    Not in our library10.1083/jcb.200404159

  258. 258.

    Yusof, A.M., N.J. Hu, A. Wlodawer, and A. Hofmann. 2005. Structural evidence for variable oligomerization of the N-terminal domain of cyclase-associated protein (CAP). Proteins. 58:255–262.

    Not in our library10.1002/prot.20314

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