The Nature of Cofilin’s Severing Mechanism by The Yale Undergraduate Research Journal

Lester | Structural Biology

The Nature of Cofilin’s Severing Mechanism By Ethan R. Lester1 1 Department of Molecular Biophysics and Biochemistry, Yale University

ABSTRACT

Cofilin plays an essential role in regulating actin filament propagation through the cytoplasmic medium. Over the years, various studies have been conducted in an attempt to better understand the complex mechanism by which cofilin promotes filament severing and de-polymerization. Here, we have compiled information obtained from these studies in order to craft a more complete and succinct description of cofilin functionality. In particular, we review the precise structural and mechanical changes associated with cofilin-binding, and the subtle ways in which some of these structural changes may be interconnected. KEYWORDS: cofilin; F-actin; microfilament; twist; tilt; D-loop; flexural rigidity; torsional rigidity; mechanical asymmetry; filament-severing; de-polymerization

INTRODUCTION

sion (Bernstein & Bamburg, 2010). Thus, the ubiquitous nature of cofilin and many other ADF proteins in normal cellular processes Cofilin is a member of the actin depolymerizing factor (ADF) pro- would pose a significant challenge to using endogenous cofilin as tein family and plays a critical role in the propagation of actin-based a drug target. microfilaments through the cytoplasmic medium (Bamburg, 1999; Carlier et al., 1997). Cellular microfilaments are composed of mo- It has long been known that ADF/cofilin binds cooperatively to nomeric actin (G-actin) that is assembled into long fibers of fila- F-actin in the cell, and catalyzes the severing and depolymerization mentous actin (F-actin). Actin filaments are known to play a crucial of these actin filaments — thereby increasing the number of barbed role in driving cell motility and other structural changes in a wide (+) ends and pointed (–) ends where newly-regenerated monomeric variety of cell types, serving multiple functions (Bravo-Cordero, actin units can either add or dissociate, respectively (Carlier et al., Magalhaes, Eddy, Hodgson, & Condeelis, 2013; Carlier et al., 1997; Enrique M. De La Cruz, 2009; Hayden, Miller, Brauweiler, 1997; Pollard & Borisy, 2003). In neural cells, for example, cofi- & Bamburg, 1993; Michelot et al., 2007; Roland, Berro, Michellin was shown to be a key regulator of dendrite morphology and, ot, Blanchoin, & Martiel, 2008). This results in a form of “steadymore recently, axon growth and regeneration (Noguchi et al., 2016; state” in which G-actin monomers are continuously “treadmilled” Tedeschi et al., 2019; Zhang et al., 2019). In light of this, it is un- through the cytoplasm from the pointed (–) ends to the barbed (+) surprising that drastic changes in cofilin activity and F-actin density ends (Carlier et al., 1997). The net result is the stochastic propagahave been shown to influence the pathogenesis of many nervous tion of these actin filaments through the cytoplasm (Roland et al., system disorders — most notably, Alzheimer’s disease (Bamburg 2008). These propagating filaments are then able to generate pi& Bernstein, 2016; Heredia et al., 2006; D. E. Kang & Woo, 2019; conewton forces that promote the formation of lamellipodia for cell Liu et al., 2019; Maloney & Bamburg, 2007; Minamide, Striegl, movement and other structural changes — such as axon- or denBoyle, Meberg, & Bamburg, 2000). Furthermore, cofilin has been drite-formation in neural cells (Bravo-Cordero et al., 2013; Enrique shown to play a role in the invasion and metastasis of many forms M. De La Cruz, 2009; Kovar & Pollard, 2004; Noguchi et al., 2016; of cancer — including breast cancer, colorectal cancer, medullary Tedeschi et al., 2019). Over the past twenty-five years, much focus thyroid carcinoma, and lung cancer (Giardino et al., 2019; Islam, has been placed on trying to better our understanding of the precise Patel, Bommareddy, Khalid, & Acevedo-Duncan, 2019; Kolegova nature of cofilin–F-actin interactions. Characterizing the exact naet al., 2019; W. Wang et al., 2006; W. G. Wang, Eddy, & Condeelis, ture of cofilin’s severing ability would pave the way for developing 2007). These examples represent only a small fraction of the disor- new pharmaceutical treatments that could instead use actin filaders in which cofilin is thought to play a role. ments as a target — perhaps by mimicking the structural impacts of cofilin-binding. Yet, our understanding of the allosteric mechanism Due to the relevance of actin filament dynamics in such a wide ar- by which cofilin severs/depolymerizes actin filaments — and of the ray of human illnesses, cofilin has been proposed as a possible ther- underlying conformational changes it induces in F-actin — continapeutic target in developing novel pharmaceutical treatments (Al- ues to evolve as new studies emerge and add to the ever-increasing hadidi, Bin Sayeed, & Shah, 2016). However, cofilin is also known complexity of cofilin functionality. Here, we present an overview to play a role in other cellular processes, including phospholipid of the various physical and structural changes that have been shown metabolism, apoptosis cascades, and mechanisms of gene expres- to contribute to cofilin’s severing and depolymerizing mechanisms.

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Lester | Structural Biology COFILIN–INDUCED MECHANICAL ASYMMETRIES The actin-severing functionality of cofilin relies on its ability to change both the local and global conformation of F-actin upon binding to it (Galkin et al., 2011; Galkin, Orlova, Lukoyanova, Wriggers, & Egelman, 2001; Galkin et al., 2003; Prochniewicz, Janson, Thomas, & De La Cruz, 2005). More specifically, it has been proposed by McCullough et al. (2008) that the main source of cofilin’s severing power arises from its ability to create two kinds of mechanical asymmetry in the filament: flexural (bending) and torsional (Enrique M. De La Cruz, 2009; McCullough, Blanchoin, Martiel, & De La Cruz, 2008). Here, we will consider individually these two types of asymmetry in an effort to concisely characterize the changes in filament mechanics that are caused by cofilin-binding. We will also take a closer look at the structural basis for each of these mechanical changes.

Figure 1. Overlaid digital images of actin (blue) and cofilactin (red) filaments undergoing thermal fluctuations in shape compared to a rigid rod (yellow) for reference. The greater bending flexibility of cofilactin (red) is evidenced by the overall increase in amplitude of the fluctuations in the image shown above. Original images obtained using fluorescence microscopy with fluorescently labeled actin. Figure borrowed from McCullough et al. (2008).

Cofilin-Binding Changes the Flexural (Bending) Rigidity of bend with higher amplitudes (to a greater extent) than bare actin F-Actin filaments (see Figure 1) . It has previously been shown using both cryogenic electron microscopy (Cryo-EM) (Orlova & Egelman, 1993) and fluorescence microscopy (McCullough et al., 2008) that cofilin-binding decreases the flexural rigidity and persistence length (LP) of actin filaments by a factor of ~4.4 (Enrique M. De La Cruz, 2009; McCullough et al., 2008; Orlova & Egelman, 1993). In a qualitative sense, the LP defines the maximum length at which a filament (or any other object) continues to behave like a rigid rod in its environment. Thus, flexural rigidity and persistence length are two distinct measures of bending flexibility. McCullough et al. (2008) showed that cofilin-bound actin filaments (also known as “cofilactin”) appear to

Figure 2. Structure of a single F-actin protofilament consisting of two protomers. Red numbers denote the four subdomains in each actin protomer, and the D-loop (lacking secondary structure) is labeled in the upper-right corner of each protomer. The hydrophobic groove between SD1 and SD3, where the D-loop interacts with the hydrophobic cleft of the next protomer, is also labeled. A space-filling model of ADP (yellow) shows the nucleotide-binding site in actin. Figure adapted from Izoré et al. (2016).

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As shown in Figure 1, the cofilactin segments bend to a much greater extent under the influence of thermal forces than the actin filaments do — indicating that the flexural rigidity of F-actin is significantly decreased when cofilin binds . This increased bending flexibility in cofilactin also corresponds to a greater critical severing angle (73 ± 7º) compared to actin — which has a critical severing angle of 54±4º (McCullough et al., 2011). Various studies have determined that the structural source of this greater bending flexibility lies in the broken longitudinal contacts between subsequent actin protomers (Bobkov et al., 2002; Enrique M. De La Cruz, 2009; Fan et al., 2013; McCullough et al., 2008). Cofilin binds to F-actin in a groove between subdomains 1 and 3 — thereby disrupting the contacts between the partially-hydrophobic DNase1-binding loop (“D-loop”; residues 38-52) in subdomain 2 (SD2) of one actin protomer and the C-terminus “hydrophobic groove” in subdomain 1 (SD1) of the subsequent protomer (Aihara & Oda, 2013; Bobkov et al., 2006; E. M. De La Cruz, 2005; Enrique M. De La Cruz, 2009; Scoville et al., 2009). More specifically, it shifts the D-Loop in SD2 away from the hydrophobic groove and closer to SD3 (see later discussion on change in tilt of the actin outer domain) (Bobkov et al., 2002; Galkin et al., 2001). Interestingly, MD simulations conducted by Fan et al. (2013) simultaneously showed an overall net decrease in the SD3-SD2 longitudinal contacts — increasing the distance between them by ~4Å. Although perhaps less influential in altering the stability of actin filaments, cofilin-binding may also shift the SD4 of one protomer farther away from SD3 of the subsequent protomer, thereby diminishing the SD4–SD3 longitudinal contact (Fan et al., 2013; Galkin et al., 2001). Overall, cofilin binding decreases the total number of longitudinal contacts from ~103 residue pairs to ~31 residue pairs, and the strength of each of these contacts is predicted to decrease by ~20-fold (Fan et al., 2013). Figure 2 (adapted from Izoré et al., 2016) shows the structure of a bare F-actin protofilament, with the four subdomains and the D-loop labeled in red and black, respectively.

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Figure 3. Graphics generated from atomistic MD simulations showing physical locations of longitudinal contacts in F-actin (left) and cofilactin (right), and the radial distributions of these contacts. The visual loss of the outer “contact ring” (turquoise) in going from actin to cofilactin corresponds to the decreased contact between SD2 (the D-loop) and SD1 (the hydrophobic groove) of the subsequent protomer. The change in intensity of the inner contact ring (denoted by the change in color from red to turquoise) corresponds to a potential decrease in SD4–SD3 contacts. The generated radial distributions of inter-protomer contacts illustrate the decrease in effective contact radius when cofilin binds to F-actin. “r” = radial distance from center of filament axis. These simulations utilized the structure of rabbit skeletal muscle F-actin with water as the solvent. Figure borrowed from Fan et al. (2013).

Furthermore, one might also argue that this broken contact between the D-loop and the hydrophobic groove is the most important factor in decreasing the flexural rigidity of the filament because the D-loop makes the highest-radius contact within the filament (Enrique M. De La Cruz, 2009; McCullough et al., 2008). Breaking this contact narrows the radial mass distribution of the filament and decreases the effective contact radius from 15.26Å in actin to 12.68Å in cofilactin, which decreases the overall stiffness of the filament by changing its geometric moment (Enrique M. De La Cruz, 2009; Fan et al., 2013; McCullough et al., 2008). Atomistic molecular dynamics (MD) simulations conducted by Fan et al. (2013) using the structure of rabbit skeletal muscle F-actin further illustrated these decreased inter-protomer contacts in cofilactin, and show how these contacts are more concentrated toward the center of the filament in cofilactin (see Figure 3) .

between the D-loop (in SD2) and hydrophobic cleft (in SD1) (Enrique M. De La Cruz, 2009).

Cofilin-Binding Changes the Torsional (Twisting) Rigidity of F-Actin Time-resolved phosphorescence anisotropy experiments on the twisting motions of labeled actin and cofilactin filaments (see Prochniewicz et al., 2005) demonstrated that cofilin-binding also decreases the torsional rigidity constant of actin filaments by a factor of ~20, which corresponds to a ~4-fold increase in the rootmean-square amplitude of the thermally-driven inter-protomer angular fluctuations (from 4.0º to 16.8º, at 25ºC) (Enrique M. De La Cruz, 2009). This large increase in angular disorder is also evidenced by the widening of the torsional angle distribution in Figure 4 (borrowed from Fan et al., 2013).

The change in the effective contact radius between SD2 and SD1 — in conjunction with the visual simulation results obtained by The widths of the distributions in Figure 4 are directly proportional Fan, et al. (2013) — suggest that the primary structural cause of the to the twisting flexibility of each type of filament. The greater width increased bending flexibility of cofilactin is this broken interaction of the cofilactin (red) probability distribution shows that cofilactin

Figure 4. Torsion angle probability distributions for bare actin (blue, dashed) and cofilactin (red) filaments, obtained from atomistic MD simulations. The broader distribution of torsion angles for cofilactin segments (red) illustrates the increased torsional variability (flexibility) of F-actin when cofilin binds to it. These simulations were conducted using the structure of rabbit skeletal muscle F-actin with water as the solvent. Figure borrowed from Fan et al. (2013).

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Lester | Structural Biology segments are much more compliant to thermal-induced twisting than bare actin filaments. For several years, there was some disagreement in the field over the relative importance of lateral inter-subunit interactions in determining the torsional rigidity of F-actin (Bobkov et al., 2006; Fan et al., 2013; McGough & Chiu, 1999). However, the MD simulations conducted by Fan et al. (2013) ultimately determined that the broken longitudinal interactions are the dominant factor in decreasing both the torsional and flexural rigidity of F-actin. In addition, mechanical simulations conducted by De La Cruz et al. (2010) and De La Cruz et al. (2015) have shown that these filament bending and twisting motions are actually coupled to one another and that this twist-bend coupling plays a substantial role in the severing ability of cofilin by amplifying the stress localized at actin-cofilactin boundaries (E. M. De la Cruz, Martiel, & Blanchoin, 2015; E. M. De La Cruz, Roland, McCullough, Blanchoin, & Martiel, 2010). These simulations also showed that up to 60% of the filament subunit elastic free energy is due to this twist-bend coupling phenomenon — which itself appears to be a direct result of the intrinsic double-protofilament helical structure of F-actin (E. M. De La Cruz et al., 2010).

Figure 5. TIRF microscopy images providing direct evidence of preferential severing at boundaries between bare (red) and cofilin-bound (green/ yellow) F-actin regions. In these experiments, a fluorescently labeled form of cofilin was genetically engineered in order to distinguish between actin and cofilactin regions. The blue arrow denotes the pointed (–) end of the actin filament, and the white arrow denotes the barbed (+) end of the filament. The orange arrow denotes the severing site, which is clearly shown to occur at the boundary between homogeneous actin (red) and cofilactin (green/yellow) regions of the filament. The second image shows the two separated filaments post-severing, approximately fifteen seconds later. Image borrowed from Suarez et al. (2011).

Actin-Severing Occurs Preferentially at Actin–Cofilactin Boundaries Due to Mechanical Heterogeneity Further studies by Enrique M. De La Cruz (2009) have also shown that severing efficiency scales with the total number of actin-cofilactin boundaries on a filament. This suggests that thermally-induced fracture events are most likely to occur at the boundaries between regions of bare actin and cofilin-decorated actin, rather than within homogenous (i.e. bare or saturated) regions of the filament (Bobkov et al., 2006; McCullough et al., 2008; Prochniewicz et al., 2005; Suarez et al., 2011). This is due to the stark change in mechanical properties and dynamics of the filaments at these junctions (E. M. De La Cruz, 2005; Enrique M. De La Cruz, 2009; McCullough et al., 2008; Prochniewicz et al., 2005). This mechanical asymmetry results in elastic stress-localization at a point immediately adjacent to the actin-cofilactin boundaries as the filaments undergo thermal twisting and bending motions (Anderson, 2005; E. M. De la Cruz et al., 2015; McCullough et al., 2008). The images in Figure 5 (borrowed from Suarez et al., 2011) were obtained using total internal reflection fluorescence microscopy (TIRF), and provide visual evidence for this preferential severing at boundaries between actin (red) and cofilactin (green/yellow) regions. Moreover, it has been shown that the effect of cofilin binding is cumulative, and as more cofilin molecules bind successive actin protomers, the filament stress localized at the (+) ends of these cofilin clusters increases dramatically (Bobkov et al., 2006; Pavlov, Muhlrad, Cooper, Wear, & Reisler, 2007). However, as the number of bound cofilin molecules increases, the number of actin-cofilactin boundaries on a filament of finite length must necessarily reach a maximum, before beginning to decrease as the filament becomes more and more saturated with cofilin. This would, in theory, result in a severing efficiency that increases initially with cofilin binding density before reaching a theoretical maximum efficiency at half-saturation, and then decreasing again as the filament becomes

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Figure 6. Experimental relationship between cofilin severing efficiency (given as a fraction of total sites on the filament) and the binding density of cofilin. The solid magenta line shows the average number of actin-cofilactin boundary sites (also expressed as a fraction of total sites) as a function of cofilin binding density, which was calculated using equilibrium binding constants from the nearest neighbor cooperative interaction model presented by Enrique M. De La Cruz, 2005 and Cao et al., 2006. Two different measures of cofilin severing efficiency are used: the filled circles show the net subunit dissociation from pointed filament ends (measured by Bobkov et al., 2006 using differential scanning calorimetry); the filled squares show the change in phase transition temperature (Tm) for a population of filaments when cofilin binds (reported by Yeoh et al., 2002). The strong correlation between the two measures of severing efficiency and the fraction of sites where an actin-cofilatin boundary exists demonstrates preferential severing at actin-cofilactin boundaries and supports the notion of a theoretical maximum severing efficiency when a filament is half-saturated with cofilin. Figure borrowed from Enrique M. De La Cruz (2009).

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Figure 7. Results from molecular simulations for three different patterns of cofilin-decoration on F-actin. (Filament length=1µm; red=actin, green=cofilactin) The top row of graphs depicts the shape of deformation that each type of actin filament would undergo when it is compressed by 1%, 10%, 20%, and 30% of its total length (black, red, purple, and blue curves, respectively). The graphs of stored elastic energy (middle row) demonstrate how cofilactin regions have higher levels of stored elastic energy at each compression level, which corresponds to a greater degree of flexibility. The graphs depicting elastic energy gradients at various points along the filament show how energy gradients reach a maximum at points within cofilactin regions that are immediately adjacent to (border) regions of bare actin (i.e. at actin-cofilactin boundaries). The white space behind the cofilactin regions is also shaded in the bottom two rows of graphs to aid in visual interpretation. Figure borrowed from De La Cruz et al. (2015). (A similar diagram can also be found in Schramm et al., 2017.)

saturated with cofilin (Andrianantoandro & Pollard, 2006; Pavlov COFILIN-INDUCED CHANGES IN THE LOCAL F-ACTIN et al., 2007). These predictions are supported by studies conducted STRUCTURE by Yeoh et al. (2002) and Bobkov et al. (2006), which are summarized in Figure 6 (borrowed from Enrique M. De La Cruz, 2009). As we previously discussed, when cofilin binds to an actin filament, longitudinal inter-subunit contacts are broken, which compromises The strong correlation shown in Figure 6 between the two measures the torsional and flexural rigidity of the filament. For many years, it of severing efficiency and the total number of boundary sites — has been known that F-actin undergoes at least two key macromoalong with the TIRF microscopy images presented by Suarez et lecular structural changes when cofilin binds: the tilt angle of the al. (2011) in Figure 5 — provide strong evidence of preferential “outer domain” (composed of SD1 and SD2) changes (see Galkin et severing at actin-cofilactin boundaries rather than within homoge- al., 2001), and the twist angle between adjacent protomers changes nous regions due to drastic changes in filament flexibility and the (see McGough et al., 1997). However, it remained unclear which localization of elastic stress at those junctions. In addition, these structural changes were responsible for the broken longitudinal results support the hypothesis of a theoretical maximum severing contact(s). Here, we present an answer to this long-held question, efficiency when the filament is half-saturated with cofilin. based on a compilation of data from previous studies conducted by others. Mechanical and thermodynamic simulations conducted by E. M. De La Cruz et al. (2015) also support the claim that filaments pref- Cofilin Disrupts SD2–SD1 (and Perhaps SD4–SD3) Longitudierentially sever at these boundaries . In particular, they showed that nal Contacts by Increasing the Local Helicity when adjacent F-actin and cofilactin regions of equal length are subjected to compressive forces, the asymmetry in the deforma- When cofilin binds to an actin filament, it alters the local twist tions of the two regions results in localization of elastic energy and angle of the filament — a change which has been shown to only a sharp increase in the total energy gradient directly adjacent to the propagates 1-2 subunits into the bare actin region (Huehn et al., boundary, but within the more flexible cofilactin region (see Figure 2018; McGough, Pope, Chiu, & Weeds, 1997). When discussing 7) (E. M. De la Cruz et al., 2015). the structure of F-actin, it is also important to note that it can be treated as either a right-handed, two-start, long-pitch double helix Moreover, the unsymmetrical deformation of the filament that is or as a left-handed, one-start, short-pitch single helix (see Figure 8) half-actin and half-cofilactin in Figure 7 provides visual evidence (Jegou & Romet-Lemonne, 2020). of this mechanical heterogeneity (i.e. differences in flexural rigidity between the two regions). Although compressive buckling forces The long-pitch torsion angle between one protomer and the longiare less relevant when discussing severing promoted by stochastic tudinally adjacent protomer (i.e. the horizontal angle between the thermal motions, these simulations nonetheless provide a very good small black dots in Figure 8) was shown to be approximately +27º description of how/where elastic energy is localized in and around in bare actin (measured by Galkin et al., 2011 using Cryo-EM). the actin-cofilactin boundaries (E. M. De la Cruz et al., 2015). This is coupled to a short-pitch torsion angle equal to –166.6º going

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Figure 8. Diagram illustrating two different ways to describe the helicity in actin filaments. Yellow labels correspond to the short-pitch, single-helix interpretation, and red labels illustrate the long-pitch, double-helix interpretation. In each case, the helical paths and connections between “subsequent protomers” according to each interpretation are labeled in the corresponding color. The length of the helical “pitch” according to each helical interpretation is also shown. The two protofilaments are shown in different shades of gray. Each sphere represents one actin monomer, and the black dots correspond to a single reference point on each monomer. Figure borrowed from Jegou and Romet-Lemonne (2020).

from one protomer to the next one above it in the single-helix (determined by Huehn et al., 2018, from a structure solved by Oda et al., 2009 using X-ray fiber diffraction). On the other hand, cofilactin filaments were shown to have a long-pitch torsion angle equal to +35.8º (measured by Tanaka et al., 2018 using Cryo-EM), and a short-pitch torsion angle equal to –162.1º (determined by Schramm et al., 2017, from a structure solved by Galkin et al., 2015 using cryo-EM). Thus, when cofilin binds to F-actin, it induces a localized increase in the helicity of the filament — including a ~9.5º increase in the long-pitch torsion angle and a ~4.5º decrease in the absolute value of the short-pitch angle (Galkin et al., 2011; Jegou & Romet-Lemonne, 2020). It might also be noted that these experimentally determined values for the short-pitch torsion angles of actin and cofilactin are comparable to the values determined from MD simulations (–165.8(±2.2)º and –163.7(±2.9)º, respectively) by Fan et al. (2013). Data from previous studies suggest that this local change in helicity causes the one (potentially two) broken inter-protomer contact(s) that increase filament flexibility. MD filament models created by Schramm et al. (2019) have shown that the ~4.5º difference in the short-pitch filament is to blame for the disrupted SD2–SD1 contact between the D-loop and the hydrophobic cleft. Interestingly, this opposes previous studies conducted by Galkin et al. (2003), which suggested that the twist did not have a direct effect on these SD2-SD1 contacts. To the best of our knowledge, it remains unclear whether or not the increase in helicity directly causes the observed net decrease in SD3-SD2 longitudinal contacts. Perhaps future simulation studies on actin filaments with constrained ends to selectively amplify the change-in-twist effect of cofilin (similar to the studies conducted by Wioland et al., 2019) might reveal the answer to this uncertainty. A low-resolution Cryo-EM structure obtained by Galkin et al. (2001) previously suggested that the other longitudinal contact between SD4 and SD3 is also disrupted by this increase in helicity. However, subsequent higher-resolution Cryo-EM structures determined by Galkin et al. (2011) and Tanaka et al. (2018) did

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not support this claim. The supposed broken contact between SD4 and SD3 was only observed in simulations conducted by Fan et al. (2013), and not in previously determined Cryo-EM structures — suggesting that perhaps these broken SD4–SD3 contacts are only transient in nature (Fan et al., 2013; Galkin et al., 2011; Tanaka et al., 2018). Carefully executed point-mutation studies may be required to determine the relative importance of these SD4-SD3 contacts in relation to the severing functionality of cofilin. In short, the increased helicity (and the increased variability in the twist angle) in cofilactin regions disrupts longitudinal SD2–SD1 and SD3–SD2 contacts (and potentially some SD4–SD3 contacts) between adjacent protomers in F-actin, and is likely one of the main contributors to cofilin’s severing ability. At this point, it is also important that we distinguish between filament “fragmentation” and filament “de-polymerization” (Pope, Gonsior, Yeoh, McGough, & Weeds, 2000). Fragmentation refers to the process of breaking off F-actin oligomers from the pointed end of a propagating microfilament, whereas de-polymerization refers to the complete de-polymerization of F-actin back into G-actin monomers — so that they can be “treadmilled” back up to the barbed end of the propagating filament to then re-polymerize. Pope et al. (2000) showed that changes in the filament twist alone were insufficient to de-polymerize F-actin (measured by the turnover rate of actin monomers), but did dramatically increased the rate of fragmentation of short oligomers from the pointed end of the filament. In fact, more recent studies conducted by Schramm et al. (2017) and Wioland et al. (2019) have shown that mechanically increasing the helicity of filaments by constraining their ends results in a ~100-fold acceleration of the fragmentation rate. These observations all suggest that the mechanism of monomeric actin turnover consists of two parallel mechanisms that go hand-in-hand; the increased twist of F-actin results in the fragmentation of the filament into short oligomers near the pointed end (which we can see visually in Figure 5), and de-polymerization is due to some other structural change — perhaps, as we will soon see, a change in the conformation of the nucleotide-binding pocket caused by a change

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Cofilin-Binding Disrupts the SD2–SD1 Longitudinal Contacts by Changing the Internal Tilt Angle Between the Inner and Outer Domain Within Each Protomer When cofilin binds to F-actin, it also induces a change in the internal structure of each actin monomer and the tilt angle between the “outer domain” of actin protomers (consisting of subdomains 1 and 2) and the “inner domain” (consisting of subdomains 3 and 4) (see Figure 9) (Fan et al., 2013; Galkin et al., 2011; Galkin et al., 2001). As depicted in Figure 9, the dihedral “tilt” angle R2–R1–R3–R4 increases from 10.27º (±3.41º) in bare actin to 25.73º (±3.71º) in cofilin-decorated actin (Fan et al., 2013). This ~15º change in the tilt of the outer domain relative to the inner domain (red arrow, Figure 9) results in the reorganization of longitudinal inter-subdomain contacts. In particular, contacts between SD2 (the D-loop) and SD1 (the hydrophobic groove) on the adjacent protomer are broken, and the SD2 is shifted closer to SD3 on the adjacent protomer (Galkin et al., 2001; Galkin, VanLoock, Orlova, & Egelman, 2002). This is one of the very same structural changes responsible for the decreased rigidity of F-actin when cofilin binds (see previous discussion on the importance of mechanical heterogeneity). We can then draw the conclusion that this “tilting” of the outer domain is another important macromolecular structural change that causes the decreased rigidity seen in cofilactin filaments. In vertebrate cofilin, it has also been shown that another important consequence of this change in the outer domain tilt angle is that it disrupts the binding pocket for an important Mg2+ stabilizing cation within the F-actin structure (H. R. Kang et al., 2014). The presence or absence of this Mg2+ cation has been shown to play a crucial role in modulating the bending and twisting mechanical properties of actin filaments (Hocky et al., 2016; H. R. Kang et al., 2014). The Mg2+ binding site is found at the SD2–SD2 interface, cradled mostly by polar/charged residues located at the base of the D-loop in SD2 (Q49, K50, D51, E57) and the inner rim of the hydrophobic cleft (E167) (see Figure 10) (Hocky et al., 2016).

Figure 9. (a) Model showing the peptide backbone of a bare actin protomer, with the four subdomains labeled R1, R2, R3, and R4. The rigid portions of each subdomain are shown in blue (res. 5-33, 80-147, 334-349), red (res. 34-39, 52-59), green (res. 148-179, 273-333), and magenta (res. 180-219, 252-262), respectively. Solid spheres of the same color denote the center of gravity (COG) for each subdomain, connected by solid gray vector lines. (b) Model showing the peptide backbone of a cofilin-bound actin protomer, viewed from a different perspective. The change in position of the subdomain 2 COG is shown, with the dihedral angle R2–R1–R3–R4 corresponding to the internal “tilt angle” within that protomer. Figure borrowed from Fan et al. (2013).

such a way that allows the hydrophobic residues in the D-loop (residues 52–56) to fit snugly into the hydrophobic cleft of SD1 (Hocky et al., 2016). However, the Mg2+ binding pocket overlaps with the cofilin binding site such that when cofilin binds, the change in tilt of the outer domain also alters the geometry of the pocket and expels the Mg2+ ion — destabilizing the filament and promoting severing (H. Kang, Bradley, Elam, & De La Cruz, 2013; H. R. Kang et al., 2014). Thus, we now have a clearer understanding of why this change in tilt promotes filament severing.

An interesting observation was that in the yeast homolog of actin, which lacks this Mg2+ binding pocket, the vertebrate cofilin is unable to sever the filaments (H. R. Kang et al., 2014). Yet, in A167E mutant yeast, where the binding pocket has been “restored,” cofilin regains its severing capabilities (H. R. Kang et al., 2014). This suggests that this Mg2+-displacement — and the broken electrostatic interactions associated with it — contributes a significant portion of This divalent cation serves as an “electrostatic linker” — adhering the fragmentation free energy. Future point-mutation studies may the SD2 of one protomer to the SD1 of the adjacent protomer in be able to determine the exact contributions of the hydrophobic

Figure 10. Structural model of F-actin depicting the location of the Mg2+ (purple sphere) binding pocket and the surrounding residues. Right image shows the same pocket from a different perspective, with the Mg2+ cation hidden. (black=SD2, orange=SD1). Figure borrowed from Hocky et al. (2016).

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Lester | Structural Biology interactions and the electrostatic interactions to the overall thermodynamic stability of the SD2–SD1 contact. It would also be interesting to compare the free energy of cofilin binding in vertebrates and yeast, to see if the difference (if there is one) between them is numerically equal to the calculated free energy contribution of the residue–Mg2+ electrostatic attractions.

filamentous actin; cryo-EM, cryogenic electron microscopy; LP, persistence length; D-Loop, DNase1-binding loop; cofilactin, cofilin-bound F-actin; COG, center of gravity; SD, subdomain; MD, molecular dynamics; TEM, transmission electron microscopy; Pi, inorganic phosphate ion; Q, glutamine; K, lysine; D, aspartate; E, glutamate; A, alanine

Inter-Subunit Tilt Modulates the Conformation of the Nucleo- Copyright for Reproduced Figures tide-Binding Pocket in F-Actin Even without decoration by cofilin, it has previously been shown that actin’s intrinsic ATPase activity results in the breaking of the same SD2–SD1 longitudinal contacts when the nucleotide binding cleft opens to release the cleaved inorganic phosphate ion (Pi) (Belmont, Orlova, Drubin, & Egelman, 1999; Galkin et al., 2003; Orlova & Egelman, 1992; Sablin et al., 2002). For that reason, Galkin et al. (2003) and Orlova et al. (2004) suggested that the tilt angle of the outer domain also regulates the conformation of the nucleotide binding pocket, and when cofilin increases this tilt angle, it effectively reverts the actin protomers back to a conformational state that favors de-polymerization. In fact, this tilt angle can be used as a measure of how widely the nucleotide binding pocket is opened — and thus a measure of Pi release (Fan et al., 2013). This conclusion is supported by further studies conducted by Suarez et al. (2011), which demonstrated that cofilin-binding accelerated Pi release from the nucleotide pocket . In this way, similar to the dual-effect of helicity changes in promoting filament fragmentation, the change in tilt between the outer domain and the inner domain within each cofilactin protomer has a two-fold effect in destabilizing actin filaments.

Figure 1 was reprinted from Journal of Molecular Biology, Vol 381, issue 3, McCullough et al., Cofilin Increases the Bending Flexibility of Actin Filaments: Implications for Severing and Cell Mechanics, 550-558, Copyright (2008), with permission from Elsevier. Figures 3, 4, and 9 were reprinted from Journal of Molecular Biology, Vol 425, issue 7, Fan et al., Molecular Origins of Cofilin-Linked Changes in Actin Filament Mechanics, 1225-1240, Copyright (2013), with permission from Elsevier. Figure 5 was reprinted from Current Biology, Vol 21, issue 10, Suarez et al., Cofilin Tunes the Nucleotide State of Actin Filaments and Severs at Bare and Decorated Segment Boundaries, 862-868, Copyright (2011), with permission from Elsevier. Figure 6 was reprinted by permission from Springer Nature: Springer, Biophysical Reviews, De La Cruz et al., Copyright (2015).

Figure 7 was reprinted from Biophysical Journal, Vol 108, issue 9, De La Cruz et al., Mechanical Heterogeneity Favors Fragmentation We can now mentally separate the two distinct structural shifts that of Strained Actin Filaments, 2270-2281, Copyright (2015), with give rise to the two parallel processes contributing to the breaking permission from Elsevier. down of microfilaments. The change in twist induced by cofilin is responsible for the increased rate of fragmentation seen at ac- Figure 8 was reprinted from Seminars in Cell & Developmental Bitin-cofilactin boundaries near the pointed (–) end of propagating fil- ology, Vol 102, Jegou & Romet-Lemonne, The many implications aments, and the change in tilt between the outer and inner domains of actin filament helicity, 65-72, Copyright (2020), with permission within each actin protomer indirectly causes the increased rate of from Elsevier. actin de-polymerization at the pointed end. Figure 10 was reprinted from J. Phys. Chem. B 2016, 120, 2, 45584567, https://pubs.acs.org/doi/10.1021/acs.jpcb.6b02741, with perCONCLUSION mission from the American Chemical Society (ACS). Further permissions related to this figure should be directed to the ACS. Here, we have succinctly summarized the many facets of cofilin’s filament-severing and filament-depolymerizing mechanism — in- Funding Sources cluding extensive discussions on the two form of mechanical/ dynamic asymmetry induced in F-actin by cofilin, the macromo- I would like to thank the Yale University Library for providing aclecular changes in local structure that occur within these cofilactin cess to the Web of Science Core Collection database and the Endregions, and the specific inter-domain longitudinal contacts that are Note X9 software. disrupted by these changes. Moreover, we have characterized the ways in which some of these structural and mechanical changes are actually intertwined in an effort to arrive at a more complete un- ACKNOWLEDGMENTS derstanding of how cofilin controls F-actin stability and dynamics. I would like to thank Dr. Julien Berro, Dr. Charles V. Sindelar, and ADDITIONAL INFORMATION Dr. Andrew Huehn from the Yale University Department of Molecular Biophysics & Biochemistry for their support and guidance Abbreviations throughout the process. ADF, actin depolymerizing factor; G-actin, globular actin; F-actin, Written for Julien Berro’s course MB&B 490: Senior Project.

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Lester | Structural Biology REFERENCES

Carlier, M. F., Laurent, V., Santolini, J., Melki, R., Didry, D., Xia, G. X., . . . Pantaloni, D. (1997). Actin depolymerizing factor (ADF/ Aihara, T., & Oda, T. (2013). Cooperative and non-cooperative cofilin) enhances the rate of filament turnover: Implication in acconformational changes of F-actin induced by cofilin. Biochemi- tin-based motility. Journal of Cell Biology, 136(6), 1307-1322. cal and Biophysical Research Communications, 435(2), 229-233. doi:10.1083/jcb.136.6.1307 doi:10.1016/j.bbrc.2013.04.076 De La Cruz, E. M. (2005). Cofilin binding to muscle and non-musAlhadidi, Q., Bin Sayeed, M. S., & Shah, Z. A. (2016). Cofilin as cle actin filaments: Isoform-dependent cooperative interactions. a Promising Therapeutic Target for Ischemic and Hemorrhagic Journal of Molecular Biology, 346(2), 557-564. doi:10.1016/j. Stroke. Translational Stroke Research, 7(1), 33-41. doi:10.1007/ jmb.2004.11.065 s12975-015-0438-2 De La Cruz, E. M. (2009). How cofilin severs an actin filament. Anderson, T. L. (2005). Fracture Mechanics: Fundamentals and Biophysical reviews, 1(2), 51-59. doi:10.1007/s12551-009-0008-5 Applications (3rd ed.). Boca Raton, FL: CRC Press. De la Cruz, E. M., Martiel, J. L., & Blanchoin, L. (2015). MeAndrianantoandro, E., & Pollard, T. D. (2006). Mechanism of actin chanical Heterogeneity Favors Fragmentation of Strained Actin filament turnover by severing and nucleation at different concentra- Filaments. Biophysical Journal, 108(9), 2270-2281. doi:10.1016/j. tions of ADF/cofilin. Molecular Cell, 24(1), 13-23. doi:10.1016/j. bpj.2015.03.058 molcel.2006.08.006 De La Cruz, E. M., Roland, J., McCullough, B. R., Blanchoin, L., Bamburg, J. R. (1999). Proteins of the ADF/cofilin family: Essential & Martiel, J. L. (2010). Origin of Twist-Bend Coupling in Actin regulators of actin dynamics. Annual Review of Cell and Develop- Filaments. Biophysical Journal, 99(6), 1852-1860. doi:10.1016/j. mental Biology, 15, 185-230. doi:10.1146/annurev.cellbio.15.1.185 bpj.2010.07.009 Bamburg, J. R., & Bernstein, B. W. (2016). Actin Dynamics and Fan, J., Saunders, M. G., Haddadian, E. J., Freed, K. F., De La Cruz, Cofilin-Actin Rods in Alzheimer Disease. Cytoskeleton, 73(9), E. M., & Voth, G. A. (2013). Molecular Origins of Cofilin-Linked 477-497. doi:10.1002/cm.21282 Changes in Actin Filament Mechanics. Journal of Molecular Biology, 425(7), 1225-1240. doi:10.1016/j.jmb.2013.01.020 Belmont, L. D., Orlova, A., Drubin, D. G., & Egelman, E. H. (1999). A change in actin conformation associated with filament instabil- Galkin, V. E., Orlova, A., Kudryashov, D. S., Solodukhin, A., Reisity after P-i release. Proceedings of the National Academy of Sci- ler, E., Schroder, G. F., & Egelman, E. H. (2011). Remodeling of ences of the United States of America, 96(1), 29-34. doi:10.1073/ actin filaments by ADF/cofilin proteins. Proceedings of the Nationpnas.96.1.29 al Academy of Sciences of the United States of America, 108(51), 20568-20572. doi:10.1073/pnas.1110109108 Bernstein, B. W., & Bamburg, J. R. (2010). ADF/Cofilin: a functional node in cell biology. Trends in Cell Biology, 20(4), 187-195. Galkin, V. E., Orlova, A., Lukoyanova, N., Wriggers, W., & Egeldoi:10.1016/j.tcb.2010.01.001 man, E. H. (2001). Actin depolymerizing factor stabilizes an existing state of F-actin and can change the tilt of F-actin subunits. Bobkov, A. A., Muhlrad, A., Kokabi, K., Vorobiev, S., Almo, S. C., Journal of Cell Biology, 153(1), 75-86. doi:10.1083/jcb.153.1.75 & Reisler, E. (2002). Structural effects of cofilin on longitudinal contacts in F-actin. Journal of Molecular Biology, 323(4), 739-750. Galkin, V. E., Orlova, A., Vanloock, M. S., Shvetsov, A., Reisler, doi:10.1016/s0022-2836(02)01008-2 E., & Egelman, E. H. (2003). ADF/cofilin use an intrinsic mode of F-actin instability to disrupt actin filaments. Journal of Cell BioloBobkov, A. A., Muhlrad, A., Pavlov, D. A., Kokabi, K., Yilmaz, gy, 163(5), 1057-1066. doi:10.1083/jcb.200308144 A., & Reisler, E. (2006). Cooperative effects of cofilin (ADF) on actin structure suggest allosteric mechanism of cofilin function. Galkin, V. E., Orlova, A., Vos, M. R., Schroder, G. F., & Egelman, Journal of Molecular Biology, 356(2), 325-334. doi:10.1016/j. E. H. (2015). Near-Atomic Resolution for One State of F-Actin. jmb.2005.11.072 Structure, 23(1), 173-182. doi:10.1016/j.str.2014.11.006 Bravo-Cordero, J. J., Magalhaes, M. A. O., Eddy, R. J., Hodgson, L., & Condeelis, J. (2013). Functions of cofilin in cell locomotion and invasion. Nature Reviews Molecular Cell Biology, 14(7), 405415. doi:10.1038/nrm3609

Galkin, V. E., VanLoock, M. S., Orlova, A., & Egelman, E. H. (2002). A new internal mode in F-actin helps explain the remarkable evolutionary conservation of actin’s sequence and structure. Current Biology, 12(7), 570-575. doi:10.1016/s0960-9822(02)00742-x

Cao, W. X., Goodarzi, J. P., & De la Cruz, E. M. (2006). Energetics and kinetics of cooperative cofilin-actin filament interactions. Journal of Molecular Biology, 361(2), 257-267. doi:10.1016/j. jmb.2006.06.019

Giardino, E., Catalano, R., Barbieri, A. M., Treppiedi, D., Mangili, F., Spada, A., . . . Peverelli, E. (2019). Cofilin is a mediator of RET-promoted medullary thyroid carcinoma cell migration, invasion and proliferation. Molecular and Cellular Endocrinology, 495, 9. doi:10.1016/j.mce.2019.110519

Spring 2021

YURJ | Vol 2.1

Lester | Structural Biology Hayden, S. M., Miller, P. S., Brauweiler, A., & Bamburg, J. R. (1993). ANALYSIS OF THE INTERACTIONS OF ACTIN DEPOLYMERIZING FACTOR WITH G-ACTIN AND F-ACTIN. Biochemistry, 32(38), 9994-10004. doi:10.1021/bi00089a015

filament barbed ends in association with formins produces piconewton forces. Proceedings of the National Academy of Sciences of the United States of America, 101(41), 14725-14730. doi:10.1073/ pnas.0405902101

Heredia, L., Helguera, P., de Olmos, S., Kedikian, G., Vigo, F. S., LaFerla, F., . . . Lorenzo, A. (2006). Phosphorylation of actin-depolymerizing factor/cofilin by LIM-kinase mediates amyloid beta-induced degeneration: A potential mechanism of neuronal dystrophy in Alzheimer’s disease. Journal of Neuroscience, 26(24), 65336542. doi:10.1523/jneurosci.5567-05.2006

Liu, T., Woo, J. A. A., Yan, Y., LePochat, P., Bukhari, M. Z., & Kang, D. E. (2019). Dual role of cofilin in APP trafficking and amyloid-beta clearance. Faseb Journal, 33(12), 14234-14247. doi:10.1096/fj.201901268R

Maloney, M. T., & Bamburg, J. R. (2007). Cofilin-mediated neurodegeneration in Alzheimer’s disease and other amyloidopathies. Hocky, G. M., Baker, J. L., Bradley, M. J., Sinitskiy, A. V., De La Molecular Neurobiology, 35(1), 21-43. doi:10.1007/bf02700622 Cruz, E. M., & Votht, G. A. (2016). Cations Stiffen Actin Filaments by Adhering a Key Structural Element to Adjacent Subunits. Jour- McCullough, B. R., Blanchoin, L., Martiel, J. L., & De La Cruz, nal of Physical Chemistry B, 120(20), 4558-4567. doi:10.1021/acs. E. M. (2008). Cofilin increases the bending flexibility of actin filajpcb.6b02741 ments: Implications for severing and cell mechanics. Journal of Molecular Biology, 381(3), 550-558. doi:10.1016/j.jmb.2008.05.055 Huehn, A., Cao, W. X., Elam, W. A., Liu, X. Q., De La Cruz, E. M., & Sindelar, C. V. (2018). The actin filament twist changes abruptly McCullough, B. R., Grintsevich, E. E., Chen, C. K., Kang, H. at boundaries between bare and cofilin-decorated segments. Jour- R., Hutchison, A. L., Henn, A., . . . De La Cruz, E. M. (2011). nal of Biological Chemistry, 293(15), 5377-5383. doi:10.1074/jbc. Cofilin-Linked Changes in Actin Filament Flexibility Promote AC118.001843 Severing. Biophysical Journal, 101(1), 151-159. doi:10.1016/j. bpj.2011.05.049 Islam, S. M. A., Patel, R., Bommareddy, R. R., Khalid, K. M., & Acevedo-Duncan, M. (2019). The modulation of actin dynamics McGough, A., & Chiu, W. (1999). ADF/cofilin weakens lateral convia atypical Protein Kinase-C activated Cofilin regulates metastasis tacts in the actin filament. Journal of Molecular Biology, 291(3), of colorectal cancer cells. Cell Adhesion & Migration, 13(1), 106- 513-519. doi:10.1006/jmbi.1999.2968 119. doi:10.1080/19336918.2018.1546513 McGough, A., Pope, B., Chiu, W., & Weeds, A. (1997). Cofilin Izore, T., Kureisaite-Ciziene, D., McLaughlin, S. H., & Lowe, J. changes the twist of F-actin: Implications for actin filament dynam(2016). Crenactin forms actin-like double helical filaments regulat- ics and cellular function. Journal of Cell Biology, 138(4), 771-781. ed by arcadin-2. Elife, 5, 18. doi:10.7554/eLife.21600 doi:10.1083/jcb.138.4.771 Jegou, A., & Romet-Lemonne, G. (2020). The many implications Michelot, A., Berro, J., Guerin, C., Boujemaa-Paterski, R., Staiger, of actin filament helicity. Seminars in Cell & Developmental Biol- C. J., Martiel, J. L., & Blanchoin, L. (2007). Actin-filament stochasogy, 102, 65-72. doi:10.1016/j.semcdb.2019.10.018 tic dynamics mediated by ADF/cofilin. Current Biology, 17(10), 825-833. doi:10.1016/j.cub.2007.04.037 Kang, D. E., & Woo, J. A. (2019). Cofilin, a Master Node Regulating Cytoskeletal Pathogenesis in Alzheimer’s Disease. Journal of Minamide, L. S., Striegl, A. M., Boyle, J. A., Meberg, P. J., & Alzheimers Disease, 72, S131-S144. doi:10.3233/jad-190585 Bamburg, J. R. (2000). Neurodegenerative stimuli induce persistent ADF/cofilin-actin rods that disrupt distal neurite function. Kang, H., Bradley, M. J., Elam, W. A., & De La Cruz, E. M. (2013). Nature Cell Biology, 2(9), 628-636. Retrieved from <Go to ISI>:// Regulation of Actin by Ion-Linked Equilibria. Biophysical Journal, WOS:000089250600015 105(12), 2621-2628. doi:10.1016/j.bpj.2013.10.032 Noguchi, J., Hayama, T., Watanabe, S., Ucar, H., Yagishita, S., Kang, H. R., Bradley, M. J., Cao, W. X., Zhou, K. F., Grintsevich, Takahashi, N., & Kasai, H. (2016). State-dependent diffusion of E. E., Michelot, A., . . . De la Cruz, E. M. (2014). Site-specific cat- actin-depolymerizing factor/cofilin underlies the enlargement and ion release drives actin filament severing by vertebrate cofilin. Pro- shrinkage of dendritic spines. Scientific Reports, 6, 9. doi:10.1038/ ceedings of the National Academy of Sciences of the United States srep32897 of America, 111(50), 17821-17826. doi:10.1073/pnas.1413397111 Oda, T., Iwasa, M., Aihara, T., Maeda, Y., & Narita, A. (2009). The Kolegova, E. S., Kakurina, G. V., Kondakova, I. V., Dobrodeev, A. nature of the globular-to fibrous-actin transition. Nature, 457(7228), Y., Kostromitskii, D. N., & Zhuikova, L. D. (2019). Adenylate Cy- 441-445. doi:10.1038/nature07685 clase-Associated Protein 1 and Cofilin in Progression of Non-Small Cell Lung Cancer. Bulletin of Experimental Biology and Medicine, Orlova, A., & Egelman, E. H. (1992). STRUCTURAL BASIS FOR 167(3), 393-395. doi:10.1007/s10517-019-04534-9 THE DESTABILIZATION OF F-ACTIN BY PHOSPHATE RELEASE FOLLOWING ATP HYDROLYSIS. Journal of Molecular Kovar, D. R., & Pollard, T. D. (2004). Insertional assembly of actin Biology, 227(4), 1043-1053. doi:10.1016/0022-2836(92)90520-t

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Lester | Structural Biology Orlova, A., & Egelman, E. H. (1993). A conformational change in Tunes the Nucleotide State of Actin Filaments and Severs at Bare the actin subunit can change the flexibility of the actin filament. and Decorated Segment Boundaries. Current Biology, 21(10), 862Journal of Molecular Biology, 232(2), 334-341. doi:10.1006/ 868. doi:10.1016/j.cub.2011.03.064 jmbi.1993.1393 Tanaka, K., Takeda, S., Mitsuoka, K., Oda, T., Kimura-Sakiyama, Orlova, A., Shvetsov, A., Galkin, V. E., Kudryashov, D. S., Ruben- C., Maeda, Y., & Narita, A. (2018). Structural basis for cofilin bindstein, P. A., Egelman, E. H., & Reisler, E. (2004). Actin-destabi- ing and actin filament disassembly. Nature Communications, 9. lizing factors disrupt filaments by means of a time reversal of po- doi:10.1038/s41467-018-04290-w lymerization. Proceedings of the National Academy of Sciences of the United States of America, 101(51), 17664-17668. doi:10.1073/ Tedeschi, A., Dupraz, S., Curcio, M., Laskowski, C. J., Schaffran, pnas.0407525102 B., Flynn, K. C., . . . Bradke, F. (2019). ADF/Cofilin-Mediated Actin Turnover Promotes Axon Regeneration in the Adult CNS. NeuPavlov, D., Muhlrad, A., Cooper, J., Wear, M., & Reisler, E. (2007). ron, 103(6), 1073-+. doi:10.1016/j.neuron.2019.07.007 Actin filament severing by cofilin. Journal of Molecular Biology, 365(5), 1350-1358. doi:10.1016/j.jmb.2006.10.102 Wang, W., Mouneimne, G., Sidani, M., Wyckoff, J., Chen, X., Makris, A., . . . Condeelis, J. S. (2006). The activity status of coPollard, T. D., & Borisy, G. G. (2003). Cellular motility driven by filin is directly related to invasion, intravasation, and metastasis assembly and disassembly of actin filaments. Cell, 112(4), 453-465. of mammary tumors. Journal of Cell Biology, 173(3), 395-404. doi:10.1016/s0092-8674(03)00120-x doi:10.1083/jcb.200510115 Pope, B. J., Gonsior, S. M., Yeoh, S., McGough, A., & Weeds, A. Wang, W. G., Eddy, R., & Condeelis, J. (2007). The cofilin pathway G. (2000). Uncoupling actin filament fragmentation by cofilin from in breast cancer invasion and metastasis. Nature Reviews Cancer, increased subunit turnover. Journal of Molecular Biology, 298(4), 7(6), 429-440. doi:10.1038/nrc2148 649-661. doi:10.1006/jmbi.2000.3688 Wioland, H., Jegou, A., & Romet-Lemonne, G. (2019). TorsionProchniewicz, E., Janson, N., Thomas, D. D., & De La Cruz, E. M. al stress generated by ADF/cofilin on cross-linked actin filaments (2005). Cofilin increases the torsional flexibility and dynamics of boosts their severing. Proceedings of the National Academy of actin filaments. Journal of Molecular Biology, 353(5), 990-1000. Sciences of the United States of America, 116(7), 2595-2602. doi:10.1016/j.jmb.2005.09.021 doi:10.1073/pnas.1812053116 Roland, J., Berro, J., Michelot, A., Blanchoin, L., & Martiel, J. L. (2008). Stochastic severing of actin filaments by actin depolymerizing factor/cofilin controls the emergence of a steady dynamical regime. Biophysical Journal, 94(6), 2082-2094. doi:10.1529/biophysj.107.121988

Yeoh, S., Pope, B., Mannherz, H. G., & Weeds, A. (2002). Determining the differences in actin binding by human ADF and cofilin. Journal of Molecular Biology, 315(4), 911-925. doi:10.1006/ jmbi.2001.5280

Zhang, X. F., Ajeti, V., Tsai, N., Fereydooni, A., Burns, W., Murrell, Sablin, E. P., Dawson, J. F., VanLoock, M. S., Spudich, J. A., Egel- M., . . . Forscher, P. (2019). Regulation of axon growth by myosin man, E. H., & Fletterick, R. J. (2002). How does ATP hydrolysis II-dependent mechanocatalysis of cofilin activity. Journal of Cell control actin’s associations? Proceedings of the National Academy Biology, 218(7), 2329-2349. doi:10.1083/jcb.201810054 of Sciences of the United States of America, 99(17), 10945-10947. doi:10.1073/pnas.152329899 Schramm, A. C., Hocky, G. M., Voth, G. A., Blanchoin, L., Martiel, J. L., & De La Cruz, E. M. (2017). Actin Filament Strain Promotes Severing and Cofilin Dissociation. Biophysical Journal, 112(12), 2624-2633. doi:10.1016/j.bpj.2017.05.016 Schramm, A. C., Hocky, G. M., Voth, G. A., Martiel, J. L., & De La Cruz, E. M. (2019). Plastic Deformation and Fragmentation of Strained Actin Filaments. Biophysical Journal, 117(3), 453-463. doi:10.1016/j.bpj.2019.06.018 Scoville, D., Stamm, J. D., Altenbach, C., Shvetsov, A., Kokabi, K., Rubenstein, P. A., . . . Reisler, E. (2009). Effects of Binding Factors on Structural Elements in F-Actin. Biochemistry, 48(2), 370-378. doi:10.1021/bi801649j Suarez, C., Roland, J., Boujemaa-Paterski, R., Kang, H., McCullough, B. R., Reymann, A. C., . . . Blanchoin, L. (2011). Cofilin

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