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Mechanical regulation of formin-dependent actin polymerization
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Mechanical regulation of formin-dependent actin polymerization Shimin Lea,b, Miao Yub, Alexander Bershadskyb,c,*, Jie Yana,b,d,* a
Department of Physics, National University of Singapore, Singapore 117542, Singapore Mechanobiology Institute, National University of Singapore, Singapore 117411, Singapore Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot 76100, Israel d Centre for Bioimaging Sciences, National University of Singapore, Singapore 117546, Singapore b c
A R T I C LE I N FO
A B S T R A C T
Keywords: Formin Actin polymerization Force Torque Single-molecule technologies
The actomyosin cytoskeleton network plays a key role in a variety of fundamental cellular processes such as cell division, migration, and cell adhesion. The functions of cytoskeleton rely on its capability to receive, generate, respond to and transmit mechanical signals throughout the cytoskeleton network within the cells and throughout the tissue via cell-extracellular matrix and cell-cell adhesions. Crucial to the cytoskeleton’s functions is actin polymerization that is regulated by many cellular factors. Among these factors, the formin family proteins, which bind the barbed end of an actin filament (F-actin), are known to be a major actin polymerization promoting factor. Mounting evidence from single-molecule mechanical manipulation experiments have suggested that formin-dependent actin polymerization is sensitively regulated by the force and torque applied to the F-actin, making the formin family an emerging mechanosensing factor that selectively promotes elongation of the F-actin under tensile forces. In this review, we will focus on the current understanding of the mechanical regulation of formin-mediated actin polymerization, the key technologies that have enabled quantification of formin-mediated actin polymerization under mechanical constraints, and future perspectives and studies on molecular mechanisms involved in the mechanosensing of actin dynamics.
1. Formin/profilin-mediated actin polymerization The actin cytoskeleton is involved in various physiological and pathological functions, such as cell migration, differentiation, embryo development, and cancer metastasis [1]. The highly dynamic organization of the actin cytoskeleton network is tightly controlled by a variety of proteins that regulate actin nucleation, polymerization, depolymerization, branching, bundling, and localization [2,3]. The factors that facilitate actin polymerization include the Ena/ VASP family proteins and the formin family proteins [2,4–10]. These factors, which can associate with the barbed end of the actin filament, interact with both F-actin and actin monomers, thereby able to promote actin polymerization by recruiting actin monomers to the F-actin barbed end in a processive manner. Among them, formin-mediated actin polymerization is particularly interesting, as the filament-associated formin is proposed to be sensitive to tensile forces and, therefore, regulates actin polymerization in a force-dependent manner [2] (Fig. 1). The FH1 and FH2 domains in the different formin isoforms are conserved across species [4]. Two formin monomers form a homodimer, which encircles an actin filament at the barbed end [11–13]
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(Fig. 1). The N-terminus of each FH2 monomer is linked to an intrinsically disordered FH1 domain containing multiple polyproline tracks that interact with the actin-binding protein profilin with an affinity of 3.4–17.8 μM [14,15]. Profilin has a high affinity to G-actin monomers (kD ∼ 0.1 μM) [16–20] and a much lower affinity to F-actin (kD ∼ 25 μM) [16–18]. The typical intracellular concentrations of profilin and G-actin are estimated to be in the range of 10−100 μM [21–23], and 10−300 μM [23,24], respectively. Due to the comparable concentrations and the high affinity interaction between profilin and Gactin, the profilin-bound G-actin (denoted as profilin/G-actin complex in this review) is generally acknowledged as the main polymerizable monomeric actin form [25]. It has been shown in vitro that profilin greatly accelerates filament assembly by formins [2,4,10,13,15,18,23,25–31]. At given G-actin concentrations, the profilin-dependent actin polymerization biphasically depends on profilin concentration, with an optimal G-actin: profilin ratio of ∼1:4 [13,29,26–31]. It has been widely discussed how the interplay among profilin, Gactin, polyproline tracks on FH1 and the F-actin barbed end results in the profilin-dependent actin polymerization mediated by formins. One view is that the profilin/G-actin complexes are bound to the polyproline
Corresponding authors at: Mechanobiology Institute, National University of Singapore, Singapore 117411, Singapore. E-mail addresses: [email protected] (A. Bershadsky), [email protected] (J. Yan).
https://doi.org/10.1016/j.semcdb.2019.11.016 Received 31 July 2019; Received in revised form 19 November 2019; Accepted 27 November 2019 1084-9521/ © 2019 Elsevier Ltd. All rights reserved.
Please cite this article as: Shimin Le, et al., Seminars in Cell and Developmental Biology, https://doi.org/10.1016/j.semcdb.2019.11.016
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Fig. 1. Formin mDia1 at a glance. (A) Illustrations of the domain map of a formin mDia1 molecule (top panel) and a mDia1 dimer attached to F-actin barbed end under tensile force (bottom panel). (B) Illustration of the mechanical gate model of force-dependent formin-mediated actin polymerization, proposed by Michael M. Kozlov and Alexander D. Bershadsky [38]. The light pink and red arrows indicate the elastic forces tending to restore the initial relative position of the formin subunits. The blue arrows indicate the external pulling forces applied to formin subunits. (C) Illustration of a “looping” model where the FH1 domain is looped to the F-actin barbed end through interaction between the profilin/G-actin complexes bound on FH1 polyproline tracks and F-actin barbed ends. Illustration is modified based on Cao et al. [32]. (D) Illustration of a local G-actin concentration enrichment model where the local G-actin or profilin/G-actin concentration is effectively increased through dynamic ternary interaction between profilin, G-actin, and FH1 polyproline tracks. Illustration is modified based on Yu et al. [31].
applied to actin polymerization by formin/profilin could result in faster recruitment of profilin/G-actin complexes to the barbed end through a combination of one-dimensional sliding diffusion on FH1 track and three-dimensional diffusion of dissociated profilin/G-actin. Unlike the juxtapositioning mechanism, the diffusion-based mechanism is not suppressed by force.
tracks on FH1, and are directly delivered to F-actin barbed ends through a ternary interaction between profilin/G-actin complexes, polyproline tracks, and F-actin [13,23,30,32]. This model requires juxtapositioning or looping between the profilin/G-actin bound polyproline tracks and F-actin barbed ends (Fig. 1C). Thus, this model implies that the farther the polyproline tracks are from the barbed end, the less effective profilin would be in promoting actin polymerization. In addition to the effects of juxtapositioning on the direct delivery of profilin/G-actin complexes to the F-actin barbed ends, it has also been proposed how the interactions between profilin, G-actin, and FH1 polyproline tracks could increase the effective local concentration of profilin/G-actin complexes, which may then increase the rate of G-actin incorporation into the F-actin barbed ends under certain conditions (Fig. 1D) [31]. The interactions can greatly enrich the local concentration of the bound profilin/G-actin complexes in the vicinity of Factin barbed ends, within a few nanometers (the size of coiled FH1 domain) [31]. Previous studies have shown that both the polyproline/ profilin interaction and the profilin/G-actin interaction are in rapid equilibrium [16,17,25]. Therefore, it is expected that the profilin/Gactin complexes dynamically bind to and dissociate from the FH1 polyproline tracks, and potentially can slide from one track to another on FH1. It has been shown that the combination of the two types of diffusion can drastically speed up the rate of target searching of transcription factor to specific DNA binding sites [33]. Similar mechanism
2. Mechanical regulation of formin-dependent actin polymerization 2.1. Force-sensing through the FH2 domain It has been shown that the FH2 homodimer associated with F-actin barbed end acts as a gate, slowing down the recruitment of actin monomers to the F-actin barbed end compared to a free F-actin barbed end [12,13]. Therefore, any physical or chemical factor that antagonizes this suppression can potentially promote the rate of forminmediated actin polymerization. Since formins can tether to the membrane and other cellular structures through various adapter proteins, formins attached to F-actin barbed ends are believed to be subject to mechanical stretching, caused by actomyosin contractility [4,26,34–37]. This raises the possibility that force may perturb FH2 conformation and mechanically regulate the rate of actin polymerization. 2
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Table 1 Summary of the experimental values of force-dependent rate of actin polymerization mediated by formin/profilin.
The experimental conditions and resulting rate values are summarized from recent reported studies by different single-molecule manipulation technologies [27–29,31,40]. The values in gray-shaded boxes are obtained from experiments in the absence of profilin. More details can be found in the corresponding experiments [27–29,31,40].
when the relative rotation between the filament and mDia1 is suppressed or inhibited. Consistently, when the relative rotation between the filament and formin (Mdia1) is allowed, the filament was shown to processively rotate in a right-handed manner with respect to the immobilized formin, during elongation [37]. On the other hand, a previous study showed that, when relative rotation between the filament and formin (Bni1p or Cdc12p) is inhibited, the growing filament buckled but did not supercoil [42]. Based on this observation, the authors proposed that formins do not stair step along the two subunits exposed on the growing barbed end. An alternative explanation was proposed that when the accumulated torque exceeds a certain level, formin may loosen its association with the F-actin and reset its orientation in order to release the torque [43]. With such rotation-sliding mechanism, the accumulated torque may not be enough to drive supercoiling of F-actin.
In this regard, it was proposed that forces applied to the forminactin attachment may mediate a conformational transition of the FH2 homodimer from a suppressed “closed” conformation to a less suppressed “open” conformation, leading to an increase in the rate of actin polymerization (Fig. 1B) [38,39]. Several groups have recently tested this 'mechanical gate' hypothesis on single formin-associated actin filaments subjected to mechanical forces using different single-molecule approaches [27–29,31,40,41] (Table 1, Fig. 2). The hypothesis was directly supported by two such studies that reported a few folds increase in formin mDia1-mediated actin polymerization rate under a few pN forces, in the absence of profilin [29,40]. However, contrasting results were observed for the effects of force on Bni1p-mediated actin polymerization [27,40]. In one study where actin was stretched using magnetic tweezers, force-dependent acceleration of formin-dependent actin polymerization in the absence of profilin was also observed for Bni1p-mediated actin polymerization in a manner similar to that observed for mDia1-mediated actin polymerization [40]. An opposite effect of force on actin polymerization rates was observed in another study that performed flow-stretching of actin filaments anchored to a supported lipid bilayer [27]. Whether such discrepancy is caused by different actin tethering geometries or other factors requires further experimental investigations.
2.3. Potential mechanosensing role of FH1 domain Formin-dependent actin polymerization is also mediated by the FH1 domain through an interaction between the polyproline tracks, profilin/G-actin complexes and F-actin barbed ends [14–20]. When formin is under force, the FH1 domain is mechanically stretched. It has been proposed that force applied to FH1 might alter its conformation and affect its interaction with profilin/G-actin complexes, resulting in mechanosensing through this domain [2]. This hypothesis was recently tested on formin mDia1-dependent actin polymerization [28,40]. In one such study, formin mDia1 tethered to the surface either through FH1 or FH2 domains did not show obvious differences in force-dependent actin polymerization rates, in the presence of profilin [28]. In another study where formin mDia1 was tethered to the surface through FH1 domain [31], a similar force-dependent fold-increase in actin polymerization rates was observed, both in the absence and presence of profilin [40].
2.2. Torque-sensing through the FH2 domain Interestingly, data obtained from magnetic tweezers experiments also revealed that the force-dependent acceleration of actin polymerization requires rotation of the FH2 dimer relative to the actin filament [31,40]. The actin filament still elongates when this rotation is not allowed; however, the polymerization rate is insensitive to the force applied to the filament. This result suggests that, besides sensing force, the mDia1 FH2 also senses the torque accumulated in the filament 3
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Fig. 2. Illustrations of single-molecule manipulation technologies implemented for studies on the mechanical regulation of formin-dependent actin polymerization. (A) Illustrations of typical flow-stretching experiments. A formin dimer is tethered to a coverslip surface through its FH1 (left panel) or FH2 domains (middle panel), or to a lipid bilayer coated coverslip surface (right panel) via specific-tag pairs (indicated by light blue circles). Force generated from flow is applied to the formin-actin system (indicated by right gray arrows). The change in filament length is measured by tracking the fluorescence labeled region of the filament (indicated by yellow colored actin). For formins anchored to a lipid bilayer, a nano-barrier with a height of ∼ 10 nm is used to trap the formin-tethered lipid within the barrier (right panel). (B) Illustrations of typical optical tweezer stretching experiments. The pointed-end region is bound with a polystyrene bead via specific-tag pairs and the barbed-end of the filament is bound by formin, with its FH1 domain further tethered to another polystyrene bead via specific-tag pairs. The two polystyrene beads (usually 1–4 μ m in diameter) are trapped by two focused laser beams, which exert tensile forces to the formin-actin tether and track changes in filament length by measuring bead-to-bead distance change. (C) Illustrations of typical magnetic tweezer stretching experiments. The pointed-end region of the filament is tethered to an inactive-myosin-coated polystyrene bead immobilized on a coverslip, and the barbed-end of the filament is bound by formin, with its FH1 domain further tethered to a superparamagnetic bead via specific-tag pairs (upper panel) or through a flexible DNA linker coated on the bead (bottom panel). A constant force is applied through a pair of magnets to the superparamagnetic bead, and the change in filament length is tracked by measuring the bead-to-bead distance change. Here, we note that the scales of the illustrations are not proportional to the dimensions of the subjects. Illustrations are modified based on previous publications [27–29,31,40].
such FH1 stabilization may depend on interactions between profilin/Gactin-bound FH1 polyproline tracks and F-actin barbed ends. This requires juxtapositioning between the FH1 polyproline tracks and F-actin barbed ends, leading us to speculate that forces applied to FH1 may disrupt this interaction and thus destabilise formins at F-actin barbed ends (Fig. 1C). Consistent with this, it has been observed that force applied to FH1 significantly decreases the lifetime of mDia1 bound to Factin barbed ends [32]. Thus, the FH1 domain may play a mechanosensing role through regulating the lifetime of formin at F-actin barbed ends, in a force-dependent manner.
Results from both studies strongly suggest that FH1 domain does not contribute significantly to the force-dependence of mDia1/profilinmediated actin polymerization rates. The formin FH1 domain is believed to extend due to pulling forces, which would cause its polyproline tracks to move farther away and prevent any juxtapositioning with F-actin barbed ends. However, such force-mediated conformational changes in FH1 does not decrease mDia1/profilin-mediated actin polymerization rates; this could be explained by the diffusion of the locally enriched profilin/G-actin complexes to F-actin barbed ends, which does not require juxtapositioning between FH1 polyproline tracks and the F-actin barbed ends, against force. On the other hand, it has been reported that profilin/FH1 interaction stabilizes formin at the F-actin barbed ends, which results in increased lifetime of formin on the actin filament [10,13,32]. However,
3. Technologies to quantify formin-dependent actin polymerization under mechanical constraints Recent progress in our understanding of the mechanosensing 4
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simultaneously measure multiple F-actin, the throughput of most optical tweezers-based experiments is significantly lower, since it can manipulate only a single F-actin at a time.
mechanisms involved in formin-dependent actin polymerization relies on the mechanical manipulation of single F-actin and the precise measurement of actin elongation rates under given mechanical constraints. The single-molecule manipulation technologies used to apply mechanical constraints to F-actin mainly include flow-stretching [27,28,32,44], optical tweezers-stretching [29,45], and magnetic tweezers-stretching [31,40] (Fig. 2). These techniques apply different types of mechanical constraints to the F-actin, which might contribute to variations in the observed effects of forces on formin-dependent actin polymerization.
3.3. Magnetic tweezers Similar to optical tweezers, magnetic tweezers are also used for single F-actin stretching studies [31,40] (Fig. 2C). The pointed-end of an F-actin is attached to an immobilized polystyrene bead on the coverslip surface through F-actin-binding proteins, such as inactive myosin. The formin-attached barbed-end of the F-actin is bound to a superparamagnetic bead through specific-tag pairs (GST/anti-GST, biotinstreptavidin, etc.) attached on the FH1 domain and the bead surface, respectively. A pair of magnets is used to apply forces to the superparamagnetic bead, which is then transmitted to the formin and the Factin. F-actin elongation is measured by tracking changes in the beadto-bead distance. Filament-surface and bead-surface interactions can also be eliminated by applying a force ∼ 10-15° above the surface. Thus, the longer the F-actin grows, the farther the F-actin and the bead are away from the surface. In typical magnetic tweezers setups, the magnetic field is perpendicular to the force direction. Therefore, the super-paramagnetic bead cannot rotate around the force axis, which in turn inhibits the rotation of the formin relative to the F-actin filament. Since F-actin polymerization requires its rotation relative to the FH2 homodimer, an elongating F-actin leads to accumulation of torque in the F-actin. It has been shown that free rotation of formin around the Factin can be achieved by introducing an additional DNA linker between the FH1 domain and the super-paramagnetic bead [31,40] (Fig. 2C). The advantages of magnetic tweezers include high spatial and temporal resolutions (similar to optical tweezers), simultaneous tracking of multiple F-actin-bound beads in the same view area (similar to flow-stretching experiments), the capability of applying both force and rotation constraints to the F-actin, and the capability of tracking molecular tethers that are longer than the linear dimension of the view area (by moving the sample stage to keep the formin-attached bead within the view area). Unlike flow-stretching experiments where force changes as F-actin elongates, or optical tweezers that require forceclamp feedback to keep the force constant, a passive, constant force can be applied over a wide force range in magnetic tweezers.
3.1. Flow-stretching An F-actin is tethered to a formin that is immobilized on a coverslip surface [28,32,44] or on a mobile, supported lipid bilayer membrane [27] through specific tag-pairs (GST/anti-GST, biotin-streptavidin, etc.), in a microfluidics channel (Fig. 2A). The level of force applied to the Factin is controlled by modulating flow speeds, and F-actin length is measured using fluorescence imaging. In such experiments, force along the filament changes as the filament elongates. While such assay has advantages of convenient force generation, easy length measurement and the capability of simultaneously tracking multiple filaments, due to close proximity between the F-actin and surface, there is a concern on the potential interaction between the F-actin and the surface that might affect the estimation of the level of force applied to the F-actin. In the case of formin immobilized on a coverslip surface, F-actin rotates in a right-handed manner. This presumably results in a negative torque applied to the F-actin due to flow drag, with the torque magnitude increasing linearly along with F-actin length growth. Therefore, while the rotation is not completely inhibited, the observed force dependence of the actin polymerization rate might be affected by the torque in the F-actin. In contrast, formins anchored to mobile lipid bilayer membranes are expected to rotate freely; leading to potentially negligible effects from negative torque caused by actin elongation. However, due to the fluidity of the lipid bilayer membrane, barriers taller than 10 nm (Fig. 2A, right panel) are needed to trap a molecule within any region of the membrane [27]. Under flow, formin-attached F-actin are pushed against the barrier, which might result in unwanted friction between the F-actin and barrier, and potentially affect actin polymerization.
4. Future perspectives 3.2. Optical tweezers 4.1. The universality of mechanical gate model among formin isoforms Optical tweezers are used to stretch a single F-actin tethered between two beads, using two focused laser beams [29] (Fig. 2B). One bead is attached to the barbed end of the F-actin through mDia1 immobilized on the bead surface, while the other bead is attached to a site near the pointed-end region through specific-tag pairs (such as the frequently used biotin-streptavidin pair). F-actin elongation is quantified by measuring bead-to-bead distance, and the force applied to the Factin is controlled by changing the trap-to-trap distance. Advantages of optical tweezers include high spatial and temporal resolutions of F-actin length and accurate force calibration, as well as the capability of applying a constant force to the F-actin through a force-clamping feedback system [46]. Since the F-actin is far from the coverslip surface, it eliminates potential friction between the F-actin and the surface. In contrast to a common view that a polystyrene bead in a focused laser beam can rotate freely, several recent studies have shown that the rotation of a trapped polystyrene bead is restrained to different extents, depending on the power of the laser [47]. Therefore, the reported forcedependent actin polymerization mediated by mDia1 using the optical tweezers might be affected by the torque in the F-actin to certain extent [29,48]. It is possible to enable free rotation of formins around the Factin by introducing an additional rotationally unconstrained DNA or peptide polymer linker between the FH1 domain and the bead, which relaxes the torque. Compared to flow-stretching experiments that can
The predictions of the mechanical gate model [38] on force-induced acceleration of formin-dependent actin polymerization has been supported by findings from recent single-molecule manipulation experiments that used mDia1 as the model system [28,29,31,40]. The size of the formin FH2 domain is highly conserved [4] among the 15 isoforms found in mammals [5]. Therefore, it is reasonable to postulate that the mechanical gate model is likely a conserved mechanism for most of the formin proteins. However, the universality of the mechanical gate hypothesis is yet to be investigated for other formin family proteins. 4.2. The torque-dependence of formin-mediated actin polymerization Recent single-molecule manipulation studies have led to new findings that suggest how formins sense not only the force, but also the torque in elongating F-actin [31,40]. This observation is built upon knowledge acquired from previous structural, simulation, and experimental studies that collectively suggest how during actin polymerization, formin tracks F-actin barbed ends and rotates around the F-actin axis. Thus, when such rotation is restrained, actin polymerization is likely to stop. However, actin polymerization was still observed to progress in a force-insensitive manner under such conditions. One explanation is that when the accumulated torque exceeds a certain level, 5
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is a well-known protein that facilitates F-actin cleavage [52–56]. Interestingly, a recent single-molecule manipulation study using optical tweezers showed that cofilin-mediated F-actin cleavage is significantly suppressed by tension (in the order of tens of pN) applied to the F-actin [56]. However, contrasting observations were reported in another single-molecule manipulation study, in which tension applied to F-actin using flow-stretching was shown to have no effect on cofilin binding and only weakly enhanced F-actin cleavage [55]. However, it is to be noted that these two studies applied two different types of mechanical constraints: while optical tweezers-stretching generates homogenouslydistributed tension, flow-stretching leads to a gradient of tension along the F-actin. Whether such differences in the applied mechanical constraints led to the contrasting results obtained in the two studies remains to be investigated. Interestingly, the second study also reported that the torque in the F-actin does not hinder cofilin binding, but dramatically enhances F-actin cleavage [55]. Despite limited number of experimental studies showing the effects of forces on F-actin destruction, it is now clear that similar to actin polymerization, F-actin destruction is also mechanically regulated. These observations can be explained by force-induced and torque changes in F-actin conformational states that influence cofilin binding and cleavage [56]. Such force- and torque-dependent change in F-actin conformation that affects F-actin binding to other factors makes F-actin itself a potential mechanosensor, as discussed in a recent review [56,57].
formin may loosen its association with the F-actin and reset its orientation in order to release the torque [43]. Such rotation-sliding mechanism would allow the progression of actin polymerization; it might also induce conformation constraints in the formin, making it incapable of force-sensing. Further studies will be needed to fully understand the torque-sensing mechanism of formins. 4.3. Torque sensing function of formins in vivo The finding from recent in vitro studies that formin also senses the torque in elongating F-actin [31,40] raises an interesting question on whether it may have mechanosensing functions in live cells as well. Inside cells, formins can potentially be tethered to a variety of cellular structures [26,36,37]. Therefore, different levels of torque may be generated at the F-actin-formin linkage, depending on whether the tethered formin is allowed to rotate or not. This, in turn, may affect the extent of force-sensing involved during formin-mediated actin polymerization. The torque generated during formin-dependent actin polymerization has been proposed as the driving force that might affect the chirality of actin cytoskeleton organization in live cells [49]. The physiological importance of the torque-sensing capability of formins warrants further investigations. 4.4. Mechanical regulation of Ena/VASP-mediated actin polymerization Besides the formin family proteins, the Ena/VASP family proteins are also known to be conserved regulators of actin polymerization, and have important roles in physiological processes such as morphogenesis, axon guidance, and endothelial barrier function, as well as in pathogenic events such as cancer cell invasion and metastasis. [6–9]. The Ena/VASP family members share three main domains: 1) the N-terminal Ena/VASP homology 1 (EVH1) domain, 2) the central proline-rich domain which binds profilin and other SH3-/WW-domain containing proteins, and 3) the C-terminal EVH2 domain that includes a G-actinbinding site (GAB), an F-actin binding site (FAB) and a tetramerization coiled-coil motif at the end of C-terminus [6,9]. Interestingly, the Ena/ VASP family proteins can also be tethered to cell membranes through the EVH1 domain that interacts with a number of adaptor proteins [6,9,50,51]. For instance, the EVH1 domain binds to the focal-adhesion proteins zyxin and vinculin, which are further linked to integrins at the cell membrane [6,9], and to lamellipodin, which contains a PI(3,4)P2binding pH domain [50,51]. The EVH1 domain also binds to the axonguidance receptor Robo that is present on axonal growth cone membranes [6,9]. The fact that EVH1 domain is tethered to the membrane while the EVH2 domain is tethered to F-actin suggests that ENA/VASPmediated actin polymerization is likely influenced by mechanical forces. In addition, torque can possibly accumulate during actin polymerization if the relative rotation between the Ena/VASP tetramer and F-actin is restrained. Therefore, it is important to investigate how these mechanical constraints regulate Ena/VASP-mediated actin polymerization in future studies.
4.6. Advanced single-molecule experimental designs The implementation of various state-of-the-art single-molecule manipulation technologies has led to a better understanding of the effects of mechanical constraints on formin-mediated F-actin polymerization [27–29,31,32,40,44,45]. Most of the current technologies involve applying controlled, external tensile forces to formin-actin systems [27–29,31,32,40,44,45]. In the case of magnetic tweezers, the relative rotation between the formin and the F-actin can also be controlled [31,40]. However, the force and torque applied cannot be controlled simultaneously in any of these current technologies. Given the recent evidence that formin is also a torque sensor [31,40] new technologies need to be developed that can simultaneously apply well-controlled force and torque to F-actin and quantify the resulting formin-dependent actin polymerization under such constraints. While it is simpler to apply well-controlled, external force to a molecule, it is much more difficult to apply controlled, external torque. In this regard, a recent study implemented a modified magnetic tweezers design that can simultaneously apply force and torque to a molecule [58]. In this design, a super-paramagnetic bead is attached to the end of a DNA molecule and a nonmagnetic polystyrene bead is attached to the side of the DNA molecule. A rotating magnetic field with a constant angular velocity is applied, causing the rotation of the superparamagnetic bead and thus the rotation of the DNA molecule and the polystyrene bead with the same angular velocity. The force can be controlled by changing the distance between the magnets and the super-paramagnetic bead, while the torque can be controlled independently by changing the angular velocity of the rotating magnetic field. This design can be adapted into a transverse magnetic tweezers set up that can apply controlled force and torque during formin-mediated actin polymerization. In this setup the F-actin can be stretched in the imaging plane, as depicted in Fig. 3, which can be further integrated with fluorescence imaging of fluorescence-labeled F-actin with micropillar arrays. Finally, the experimental throughput can be increased by tracking multiple tethers in the same view area at the same time. Such integrated, multiplexing single-molecule manipulation and imaging technology has a great potential to reveal new insights into the molecular mechanisms underlying formin-dependent actin polymerization under mechanical constraints.
4.5. Mechanical regulation of F-actin destruction The mechanical regulation of formin-dependent F-actin polymerization has been well-studied; however, the mechanical principles governing F-actin destruction is not so well investigated [52–56]. The most common way for F-actin destruction is through depolymerization, where actin monomers dissociate from the F-actin ends. A recent study using single F-actin flow-stretching revealed that mDia1 remains associated at the barbed end of a depolymerizing F-actin, and the rate of depolymerization is significantly suppressed when the force applied to the F-actin is increased from 0 pN to 3 pN [28,32]. This observation suggests that force negatively regulates formin-mediated actin depolymerization. A second way of F-actin destruction is through F-actin filament cleavage, where an F-actin breaks into shorter fragments. Cofilin 6
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Fig. 3. Advanced single-molecule manipulation design for probing the torque and force effects on formin-mediated actin polymerization. (A) Illustrations of experimental designs to investigate force-dependent formin-mediated actin polymerization in a torque-free condition. In the sample channel, micro-pillar arrays (top left inset) are coupled with inactive myosin-coated non-magnetic polystyrene bead at pillar top. The pointed-end region of an actin filament is bound with the bead at pillar top. The barbed-end of the filament is bound with formin FH2 ring, while the FH1 domains are tethered to a superparamagnetic bead. A pair of magnets are used to generate forces along the filament, which magnetizes the bead in the direction perpendicular to the force direction and thus inhibits the rotation of the bead. A DNA linker (top panel) or a lipid bilayer (bottom panel) is added between the FH1 domains and the bead to enable the free rotation of formin FH2 ring around the actin filament during actin polymerization. When the FH1 domains are directly tethered to the superparamagnetic bead, rotation of formin around the actin filament can also be enabled using a cylindrically symmetric cone-shaped magnet (middle panel). In addition, the use of pillar array allows stretching of the filament in the focal plane, such that the elongating filament can be simultaneously visualized using fluorescence microscopy. (B) Illustration of an advanced magnetic tweezer design to control force and torque during formin-mediated actin polymerization. In the design, a pair of magnets is rotated with a constant angular velocity, which creates a rotating magnetic field that forces the side-attached super-paramagnetic bead and the filament to rotate with the same angular velocity. It further causes the rotation of a polystyrene bead attached to the FH1 domains of the formin dimer (bound to F-actin barbed end) with the same angular velocity. A DNA linker is used to make the polystyrene bead rotation unconstrained. The rotation drag to the polystyrene bead creates a torque at the formin-actin interface. The force can be controlled by changing the distance between the magnets and the superparamagnetic bead, while the torque can be controlled independently by changing the angular velocity of the rotating magnetic field. [5] A. Schonichen, M. Geyer, Fifteen formins for an actin filament: a molecular view on the regulation of human formins, Biochim. Biophys. Acta 1803 (2) (2010) 152–163. [6] J.E. Bear, F.B. Gertler, Ena/VASP: towards resolving a pointed controversy at the barbed end, J. Cell. Sci. 122 (Pt 12) (2009) 1947–1953. [7] D. Breitsprecher, A.K. Kiesewetter, J. Linkner, M. Vinzenz, T.E. Stradal, J.V. Small, U. Curth, R.B. Dickinson, J. Faix, Molecular mechanism of Ena/VASP-mediated actin-filament elongation, EMBO J. 30 (3) (2011) 456–467. [8] S.D. Hansen, R.D. Mullins, VASP is a processive actin polymerase that requires monomeric actin for barbed end association, J. Cell Biol. 191 (3) (2010) 571–584. [9] M. Krause, E.W. Dent, J.E. Bear, J.J. Loureiro, F.B. Gertler, Ena/VASP proteins: regulators of the actin cytoskeleton and cell migration, Annu. Rev. Cell Dev. Biol. 19 (2003) 541–564. [10] S. Romero, C. Le Clainche, D. Didry, C. Egile, D. Pantaloni, M.F. Carlier, Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis, Cell 119 (3) (2004) 419–429. [11] Y. Xu, J.B. Moseley, I. Sagot, F. Poy, D. Pellman, B.L. Goode, M.J. Eck, Crystal structures of a Formin Homology-2 domain reveal a tethered dimer architecture, Cell 116 (5) (2004) 711–723. [12] T. Otomo, D.R. Tomchick, C. Otomo, S.C. Panchal, M. Machius, M.K. Rosen, Structural basis of actin filament nucleation and processive capping by a formin homology 2 domain, Nature 433 (7025) (2005) 488–494. [13] D. Vavylonis, D.R. Kovar, B. O’Shaughnessy, T.D. Pollard, Model of formin-associated actin filament elongation, Mol. Cell 21 (4) (2006) 455–466. [14] P. Kursula, I. Kursula, M. Massimi, Y.H. Song, J. Downer, W.A. Stanley, W. Witke,
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Contents lists available at ScienceDirect
Seminars in Cell & Developmental Biology journal homepage: www.elsevier.com/locate/semcdb
Mechanical regulation of formin-dependent actin polymerization Shimin Lea,b, Miao Yub, Alexander Bershadskyb,c,*, Jie Yana,b,d,* a
Department of Physics, National University of Singapore, Singapore 117542, Singapore Mechanobiology Institute, National University of Singapore, Singapore 117411, Singapore Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot 76100, Israel d Centre for Bioimaging Sciences, National University of Singapore, Singapore 117546, Singapore b c
A R T I C LE I N FO
A B S T R A C T
Keywords: Formin Actin polymerization Force Torque Single-molecule technologies
The actomyosin cytoskeleton network plays a key role in a variety of fundamental cellular processes such as cell division, migration, and cell adhesion. The functions of cytoskeleton rely on its capability to receive, generate, respond to and transmit mechanical signals throughout the cytoskeleton network within the cells and throughout the tissue via cell-extracellular matrix and cell-cell adhesions. Crucial to the cytoskeleton’s functions is actin polymerization that is regulated by many cellular factors. Among these factors, the formin family proteins, which bind the barbed end of an actin filament (F-actin), are known to be a major actin polymerization promoting factor. Mounting evidence from single-molecule mechanical manipulation experiments have suggested that formin-dependent actin polymerization is sensitively regulated by the force and torque applied to the F-actin, making the formin family an emerging mechanosensing factor that selectively promotes elongation of the F-actin under tensile forces. In this review, we will focus on the current understanding of the mechanical regulation of formin-mediated actin polymerization, the key technologies that have enabled quantification of formin-mediated actin polymerization under mechanical constraints, and future perspectives and studies on molecular mechanisms involved in the mechanosensing of actin dynamics.
1. Formin/profilin-mediated actin polymerization The actin cytoskeleton is involved in various physiological and pathological functions, such as cell migration, differentiation, embryo development, and cancer metastasis [1]. The highly dynamic organization of the actin cytoskeleton network is tightly controlled by a variety of proteins that regulate actin nucleation, polymerization, depolymerization, branching, bundling, and localization [2,3]. The factors that facilitate actin polymerization include the Ena/ VASP family proteins and the formin family proteins [2,4–10]. These factors, which can associate with the barbed end of the actin filament, interact with both F-actin and actin monomers, thereby able to promote actin polymerization by recruiting actin monomers to the F-actin barbed end in a processive manner. Among them, formin-mediated actin polymerization is particularly interesting, as the filament-associated formin is proposed to be sensitive to tensile forces and, therefore, regulates actin polymerization in a force-dependent manner [2] (Fig. 1). The FH1 and FH2 domains in the different formin isoforms are conserved across species [4]. Two formin monomers form a homodimer, which encircles an actin filament at the barbed end [11–13]
⁎
(Fig. 1). The N-terminus of each FH2 monomer is linked to an intrinsically disordered FH1 domain containing multiple polyproline tracks that interact with the actin-binding protein profilin with an affinity of 3.4–17.8 μM [14,15]. Profilin has a high affinity to G-actin monomers (kD ∼ 0.1 μM) [16–20] and a much lower affinity to F-actin (kD ∼ 25 μM) [16–18]. The typical intracellular concentrations of profilin and G-actin are estimated to be in the range of 10−100 μM [21–23], and 10−300 μM [23,24], respectively. Due to the comparable concentrations and the high affinity interaction between profilin and Gactin, the profilin-bound G-actin (denoted as profilin/G-actin complex in this review) is generally acknowledged as the main polymerizable monomeric actin form [25]. It has been shown in vitro that profilin greatly accelerates filament assembly by formins [2,4,10,13,15,18,23,25–31]. At given G-actin concentrations, the profilin-dependent actin polymerization biphasically depends on profilin concentration, with an optimal G-actin: profilin ratio of ∼1:4 [13,29,26–31]. It has been widely discussed how the interplay among profilin, Gactin, polyproline tracks on FH1 and the F-actin barbed end results in the profilin-dependent actin polymerization mediated by formins. One view is that the profilin/G-actin complexes are bound to the polyproline
Corresponding authors at: Mechanobiology Institute, National University of Singapore, Singapore 117411, Singapore. E-mail addresses: [email protected] (A. Bershadsky), [email protected] (J. Yan).
https://doi.org/10.1016/j.semcdb.2019.11.016 Received 31 July 2019; Received in revised form 19 November 2019; Accepted 27 November 2019 1084-9521/ © 2019 Elsevier Ltd. All rights reserved.
Please cite this article as: Shimin Le, et al., Seminars in Cell and Developmental Biology, https://doi.org/10.1016/j.semcdb.2019.11.016
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Fig. 1. Formin mDia1 at a glance. (A) Illustrations of the domain map of a formin mDia1 molecule (top panel) and a mDia1 dimer attached to F-actin barbed end under tensile force (bottom panel). (B) Illustration of the mechanical gate model of force-dependent formin-mediated actin polymerization, proposed by Michael M. Kozlov and Alexander D. Bershadsky [38]. The light pink and red arrows indicate the elastic forces tending to restore the initial relative position of the formin subunits. The blue arrows indicate the external pulling forces applied to formin subunits. (C) Illustration of a “looping” model where the FH1 domain is looped to the F-actin barbed end through interaction between the profilin/G-actin complexes bound on FH1 polyproline tracks and F-actin barbed ends. Illustration is modified based on Cao et al. [32]. (D) Illustration of a local G-actin concentration enrichment model where the local G-actin or profilin/G-actin concentration is effectively increased through dynamic ternary interaction between profilin, G-actin, and FH1 polyproline tracks. Illustration is modified based on Yu et al. [31].
applied to actin polymerization by formin/profilin could result in faster recruitment of profilin/G-actin complexes to the barbed end through a combination of one-dimensional sliding diffusion on FH1 track and three-dimensional diffusion of dissociated profilin/G-actin. Unlike the juxtapositioning mechanism, the diffusion-based mechanism is not suppressed by force.
tracks on FH1, and are directly delivered to F-actin barbed ends through a ternary interaction between profilin/G-actin complexes, polyproline tracks, and F-actin [13,23,30,32]. This model requires juxtapositioning or looping between the profilin/G-actin bound polyproline tracks and F-actin barbed ends (Fig. 1C). Thus, this model implies that the farther the polyproline tracks are from the barbed end, the less effective profilin would be in promoting actin polymerization. In addition to the effects of juxtapositioning on the direct delivery of profilin/G-actin complexes to the F-actin barbed ends, it has also been proposed how the interactions between profilin, G-actin, and FH1 polyproline tracks could increase the effective local concentration of profilin/G-actin complexes, which may then increase the rate of G-actin incorporation into the F-actin barbed ends under certain conditions (Fig. 1D) [31]. The interactions can greatly enrich the local concentration of the bound profilin/G-actin complexes in the vicinity of Factin barbed ends, within a few nanometers (the size of coiled FH1 domain) [31]. Previous studies have shown that both the polyproline/ profilin interaction and the profilin/G-actin interaction are in rapid equilibrium [16,17,25]. Therefore, it is expected that the profilin/Gactin complexes dynamically bind to and dissociate from the FH1 polyproline tracks, and potentially can slide from one track to another on FH1. It has been shown that the combination of the two types of diffusion can drastically speed up the rate of target searching of transcription factor to specific DNA binding sites [33]. Similar mechanism
2. Mechanical regulation of formin-dependent actin polymerization 2.1. Force-sensing through the FH2 domain It has been shown that the FH2 homodimer associated with F-actin barbed end acts as a gate, slowing down the recruitment of actin monomers to the F-actin barbed end compared to a free F-actin barbed end [12,13]. Therefore, any physical or chemical factor that antagonizes this suppression can potentially promote the rate of forminmediated actin polymerization. Since formins can tether to the membrane and other cellular structures through various adapter proteins, formins attached to F-actin barbed ends are believed to be subject to mechanical stretching, caused by actomyosin contractility [4,26,34–37]. This raises the possibility that force may perturb FH2 conformation and mechanically regulate the rate of actin polymerization. 2
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Table 1 Summary of the experimental values of force-dependent rate of actin polymerization mediated by formin/profilin.
The experimental conditions and resulting rate values are summarized from recent reported studies by different single-molecule manipulation technologies [27–29,31,40]. The values in gray-shaded boxes are obtained from experiments in the absence of profilin. More details can be found in the corresponding experiments [27–29,31,40].
when the relative rotation between the filament and mDia1 is suppressed or inhibited. Consistently, when the relative rotation between the filament and formin (Mdia1) is allowed, the filament was shown to processively rotate in a right-handed manner with respect to the immobilized formin, during elongation [37]. On the other hand, a previous study showed that, when relative rotation between the filament and formin (Bni1p or Cdc12p) is inhibited, the growing filament buckled but did not supercoil [42]. Based on this observation, the authors proposed that formins do not stair step along the two subunits exposed on the growing barbed end. An alternative explanation was proposed that when the accumulated torque exceeds a certain level, formin may loosen its association with the F-actin and reset its orientation in order to release the torque [43]. With such rotation-sliding mechanism, the accumulated torque may not be enough to drive supercoiling of F-actin.
In this regard, it was proposed that forces applied to the forminactin attachment may mediate a conformational transition of the FH2 homodimer from a suppressed “closed” conformation to a less suppressed “open” conformation, leading to an increase in the rate of actin polymerization (Fig. 1B) [38,39]. Several groups have recently tested this 'mechanical gate' hypothesis on single formin-associated actin filaments subjected to mechanical forces using different single-molecule approaches [27–29,31,40,41] (Table 1, Fig. 2). The hypothesis was directly supported by two such studies that reported a few folds increase in formin mDia1-mediated actin polymerization rate under a few pN forces, in the absence of profilin [29,40]. However, contrasting results were observed for the effects of force on Bni1p-mediated actin polymerization [27,40]. In one study where actin was stretched using magnetic tweezers, force-dependent acceleration of formin-dependent actin polymerization in the absence of profilin was also observed for Bni1p-mediated actin polymerization in a manner similar to that observed for mDia1-mediated actin polymerization [40]. An opposite effect of force on actin polymerization rates was observed in another study that performed flow-stretching of actin filaments anchored to a supported lipid bilayer [27]. Whether such discrepancy is caused by different actin tethering geometries or other factors requires further experimental investigations.
2.3. Potential mechanosensing role of FH1 domain Formin-dependent actin polymerization is also mediated by the FH1 domain through an interaction between the polyproline tracks, profilin/G-actin complexes and F-actin barbed ends [14–20]. When formin is under force, the FH1 domain is mechanically stretched. It has been proposed that force applied to FH1 might alter its conformation and affect its interaction with profilin/G-actin complexes, resulting in mechanosensing through this domain [2]. This hypothesis was recently tested on formin mDia1-dependent actin polymerization [28,40]. In one such study, formin mDia1 tethered to the surface either through FH1 or FH2 domains did not show obvious differences in force-dependent actin polymerization rates, in the presence of profilin [28]. In another study where formin mDia1 was tethered to the surface through FH1 domain [31], a similar force-dependent fold-increase in actin polymerization rates was observed, both in the absence and presence of profilin [40].
2.2. Torque-sensing through the FH2 domain Interestingly, data obtained from magnetic tweezers experiments also revealed that the force-dependent acceleration of actin polymerization requires rotation of the FH2 dimer relative to the actin filament [31,40]. The actin filament still elongates when this rotation is not allowed; however, the polymerization rate is insensitive to the force applied to the filament. This result suggests that, besides sensing force, the mDia1 FH2 also senses the torque accumulated in the filament 3
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Fig. 2. Illustrations of single-molecule manipulation technologies implemented for studies on the mechanical regulation of formin-dependent actin polymerization. (A) Illustrations of typical flow-stretching experiments. A formin dimer is tethered to a coverslip surface through its FH1 (left panel) or FH2 domains (middle panel), or to a lipid bilayer coated coverslip surface (right panel) via specific-tag pairs (indicated by light blue circles). Force generated from flow is applied to the formin-actin system (indicated by right gray arrows). The change in filament length is measured by tracking the fluorescence labeled region of the filament (indicated by yellow colored actin). For formins anchored to a lipid bilayer, a nano-barrier with a height of ∼ 10 nm is used to trap the formin-tethered lipid within the barrier (right panel). (B) Illustrations of typical optical tweezer stretching experiments. The pointed-end region is bound with a polystyrene bead via specific-tag pairs and the barbed-end of the filament is bound by formin, with its FH1 domain further tethered to another polystyrene bead via specific-tag pairs. The two polystyrene beads (usually 1–4 μ m in diameter) are trapped by two focused laser beams, which exert tensile forces to the formin-actin tether and track changes in filament length by measuring bead-to-bead distance change. (C) Illustrations of typical magnetic tweezer stretching experiments. The pointed-end region of the filament is tethered to an inactive-myosin-coated polystyrene bead immobilized on a coverslip, and the barbed-end of the filament is bound by formin, with its FH1 domain further tethered to a superparamagnetic bead via specific-tag pairs (upper panel) or through a flexible DNA linker coated on the bead (bottom panel). A constant force is applied through a pair of magnets to the superparamagnetic bead, and the change in filament length is tracked by measuring the bead-to-bead distance change. Here, we note that the scales of the illustrations are not proportional to the dimensions of the subjects. Illustrations are modified based on previous publications [27–29,31,40].
such FH1 stabilization may depend on interactions between profilin/Gactin-bound FH1 polyproline tracks and F-actin barbed ends. This requires juxtapositioning between the FH1 polyproline tracks and F-actin barbed ends, leading us to speculate that forces applied to FH1 may disrupt this interaction and thus destabilise formins at F-actin barbed ends (Fig. 1C). Consistent with this, it has been observed that force applied to FH1 significantly decreases the lifetime of mDia1 bound to Factin barbed ends [32]. Thus, the FH1 domain may play a mechanosensing role through regulating the lifetime of formin at F-actin barbed ends, in a force-dependent manner.
Results from both studies strongly suggest that FH1 domain does not contribute significantly to the force-dependence of mDia1/profilinmediated actin polymerization rates. The formin FH1 domain is believed to extend due to pulling forces, which would cause its polyproline tracks to move farther away and prevent any juxtapositioning with F-actin barbed ends. However, such force-mediated conformational changes in FH1 does not decrease mDia1/profilin-mediated actin polymerization rates; this could be explained by the diffusion of the locally enriched profilin/G-actin complexes to F-actin barbed ends, which does not require juxtapositioning between FH1 polyproline tracks and the F-actin barbed ends, against force. On the other hand, it has been reported that profilin/FH1 interaction stabilizes formin at the F-actin barbed ends, which results in increased lifetime of formin on the actin filament [10,13,32]. However,
3. Technologies to quantify formin-dependent actin polymerization under mechanical constraints Recent progress in our understanding of the mechanosensing 4
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simultaneously measure multiple F-actin, the throughput of most optical tweezers-based experiments is significantly lower, since it can manipulate only a single F-actin at a time.
mechanisms involved in formin-dependent actin polymerization relies on the mechanical manipulation of single F-actin and the precise measurement of actin elongation rates under given mechanical constraints. The single-molecule manipulation technologies used to apply mechanical constraints to F-actin mainly include flow-stretching [27,28,32,44], optical tweezers-stretching [29,45], and magnetic tweezers-stretching [31,40] (Fig. 2). These techniques apply different types of mechanical constraints to the F-actin, which might contribute to variations in the observed effects of forces on formin-dependent actin polymerization.
3.3. Magnetic tweezers Similar to optical tweezers, magnetic tweezers are also used for single F-actin stretching studies [31,40] (Fig. 2C). The pointed-end of an F-actin is attached to an immobilized polystyrene bead on the coverslip surface through F-actin-binding proteins, such as inactive myosin. The formin-attached barbed-end of the F-actin is bound to a superparamagnetic bead through specific-tag pairs (GST/anti-GST, biotinstreptavidin, etc.) attached on the FH1 domain and the bead surface, respectively. A pair of magnets is used to apply forces to the superparamagnetic bead, which is then transmitted to the formin and the Factin. F-actin elongation is measured by tracking changes in the beadto-bead distance. Filament-surface and bead-surface interactions can also be eliminated by applying a force ∼ 10-15° above the surface. Thus, the longer the F-actin grows, the farther the F-actin and the bead are away from the surface. In typical magnetic tweezers setups, the magnetic field is perpendicular to the force direction. Therefore, the super-paramagnetic bead cannot rotate around the force axis, which in turn inhibits the rotation of the formin relative to the F-actin filament. Since F-actin polymerization requires its rotation relative to the FH2 homodimer, an elongating F-actin leads to accumulation of torque in the F-actin. It has been shown that free rotation of formin around the Factin can be achieved by introducing an additional DNA linker between the FH1 domain and the super-paramagnetic bead [31,40] (Fig. 2C). The advantages of magnetic tweezers include high spatial and temporal resolutions (similar to optical tweezers), simultaneous tracking of multiple F-actin-bound beads in the same view area (similar to flow-stretching experiments), the capability of applying both force and rotation constraints to the F-actin, and the capability of tracking molecular tethers that are longer than the linear dimension of the view area (by moving the sample stage to keep the formin-attached bead within the view area). Unlike flow-stretching experiments where force changes as F-actin elongates, or optical tweezers that require forceclamp feedback to keep the force constant, a passive, constant force can be applied over a wide force range in magnetic tweezers.
3.1. Flow-stretching An F-actin is tethered to a formin that is immobilized on a coverslip surface [28,32,44] or on a mobile, supported lipid bilayer membrane [27] through specific tag-pairs (GST/anti-GST, biotin-streptavidin, etc.), in a microfluidics channel (Fig. 2A). The level of force applied to the Factin is controlled by modulating flow speeds, and F-actin length is measured using fluorescence imaging. In such experiments, force along the filament changes as the filament elongates. While such assay has advantages of convenient force generation, easy length measurement and the capability of simultaneously tracking multiple filaments, due to close proximity between the F-actin and surface, there is a concern on the potential interaction between the F-actin and the surface that might affect the estimation of the level of force applied to the F-actin. In the case of formin immobilized on a coverslip surface, F-actin rotates in a right-handed manner. This presumably results in a negative torque applied to the F-actin due to flow drag, with the torque magnitude increasing linearly along with F-actin length growth. Therefore, while the rotation is not completely inhibited, the observed force dependence of the actin polymerization rate might be affected by the torque in the F-actin. In contrast, formins anchored to mobile lipid bilayer membranes are expected to rotate freely; leading to potentially negligible effects from negative torque caused by actin elongation. However, due to the fluidity of the lipid bilayer membrane, barriers taller than 10 nm (Fig. 2A, right panel) are needed to trap a molecule within any region of the membrane [27]. Under flow, formin-attached F-actin are pushed against the barrier, which might result in unwanted friction between the F-actin and barrier, and potentially affect actin polymerization.
4. Future perspectives 3.2. Optical tweezers 4.1. The universality of mechanical gate model among formin isoforms Optical tweezers are used to stretch a single F-actin tethered between two beads, using two focused laser beams [29] (Fig. 2B). One bead is attached to the barbed end of the F-actin through mDia1 immobilized on the bead surface, while the other bead is attached to a site near the pointed-end region through specific-tag pairs (such as the frequently used biotin-streptavidin pair). F-actin elongation is quantified by measuring bead-to-bead distance, and the force applied to the Factin is controlled by changing the trap-to-trap distance. Advantages of optical tweezers include high spatial and temporal resolutions of F-actin length and accurate force calibration, as well as the capability of applying a constant force to the F-actin through a force-clamping feedback system [46]. Since the F-actin is far from the coverslip surface, it eliminates potential friction between the F-actin and the surface. In contrast to a common view that a polystyrene bead in a focused laser beam can rotate freely, several recent studies have shown that the rotation of a trapped polystyrene bead is restrained to different extents, depending on the power of the laser [47]. Therefore, the reported forcedependent actin polymerization mediated by mDia1 using the optical tweezers might be affected by the torque in the F-actin to certain extent [29,48]. It is possible to enable free rotation of formins around the Factin by introducing an additional rotationally unconstrained DNA or peptide polymer linker between the FH1 domain and the bead, which relaxes the torque. Compared to flow-stretching experiments that can
The predictions of the mechanical gate model [38] on force-induced acceleration of formin-dependent actin polymerization has been supported by findings from recent single-molecule manipulation experiments that used mDia1 as the model system [28,29,31,40]. The size of the formin FH2 domain is highly conserved [4] among the 15 isoforms found in mammals [5]. Therefore, it is reasonable to postulate that the mechanical gate model is likely a conserved mechanism for most of the formin proteins. However, the universality of the mechanical gate hypothesis is yet to be investigated for other formin family proteins. 4.2. The torque-dependence of formin-mediated actin polymerization Recent single-molecule manipulation studies have led to new findings that suggest how formins sense not only the force, but also the torque in elongating F-actin [31,40]. This observation is built upon knowledge acquired from previous structural, simulation, and experimental studies that collectively suggest how during actin polymerization, formin tracks F-actin barbed ends and rotates around the F-actin axis. Thus, when such rotation is restrained, actin polymerization is likely to stop. However, actin polymerization was still observed to progress in a force-insensitive manner under such conditions. One explanation is that when the accumulated torque exceeds a certain level, 5
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is a well-known protein that facilitates F-actin cleavage [52–56]. Interestingly, a recent single-molecule manipulation study using optical tweezers showed that cofilin-mediated F-actin cleavage is significantly suppressed by tension (in the order of tens of pN) applied to the F-actin [56]. However, contrasting observations were reported in another single-molecule manipulation study, in which tension applied to F-actin using flow-stretching was shown to have no effect on cofilin binding and only weakly enhanced F-actin cleavage [55]. However, it is to be noted that these two studies applied two different types of mechanical constraints: while optical tweezers-stretching generates homogenouslydistributed tension, flow-stretching leads to a gradient of tension along the F-actin. Whether such differences in the applied mechanical constraints led to the contrasting results obtained in the two studies remains to be investigated. Interestingly, the second study also reported that the torque in the F-actin does not hinder cofilin binding, but dramatically enhances F-actin cleavage [55]. Despite limited number of experimental studies showing the effects of forces on F-actin destruction, it is now clear that similar to actin polymerization, F-actin destruction is also mechanically regulated. These observations can be explained by force-induced and torque changes in F-actin conformational states that influence cofilin binding and cleavage [56]. Such force- and torque-dependent change in F-actin conformation that affects F-actin binding to other factors makes F-actin itself a potential mechanosensor, as discussed in a recent review [56,57].
formin may loosen its association with the F-actin and reset its orientation in order to release the torque [43]. Such rotation-sliding mechanism would allow the progression of actin polymerization; it might also induce conformation constraints in the formin, making it incapable of force-sensing. Further studies will be needed to fully understand the torque-sensing mechanism of formins. 4.3. Torque sensing function of formins in vivo The finding from recent in vitro studies that formin also senses the torque in elongating F-actin [31,40] raises an interesting question on whether it may have mechanosensing functions in live cells as well. Inside cells, formins can potentially be tethered to a variety of cellular structures [26,36,37]. Therefore, different levels of torque may be generated at the F-actin-formin linkage, depending on whether the tethered formin is allowed to rotate or not. This, in turn, may affect the extent of force-sensing involved during formin-mediated actin polymerization. The torque generated during formin-dependent actin polymerization has been proposed as the driving force that might affect the chirality of actin cytoskeleton organization in live cells [49]. The physiological importance of the torque-sensing capability of formins warrants further investigations. 4.4. Mechanical regulation of Ena/VASP-mediated actin polymerization Besides the formin family proteins, the Ena/VASP family proteins are also known to be conserved regulators of actin polymerization, and have important roles in physiological processes such as morphogenesis, axon guidance, and endothelial barrier function, as well as in pathogenic events such as cancer cell invasion and metastasis. [6–9]. The Ena/VASP family members share three main domains: 1) the N-terminal Ena/VASP homology 1 (EVH1) domain, 2) the central proline-rich domain which binds profilin and other SH3-/WW-domain containing proteins, and 3) the C-terminal EVH2 domain that includes a G-actinbinding site (GAB), an F-actin binding site (FAB) and a tetramerization coiled-coil motif at the end of C-terminus [6,9]. Interestingly, the Ena/ VASP family proteins can also be tethered to cell membranes through the EVH1 domain that interacts with a number of adaptor proteins [6,9,50,51]. For instance, the EVH1 domain binds to the focal-adhesion proteins zyxin and vinculin, which are further linked to integrins at the cell membrane [6,9], and to lamellipodin, which contains a PI(3,4)P2binding pH domain [50,51]. The EVH1 domain also binds to the axonguidance receptor Robo that is present on axonal growth cone membranes [6,9]. The fact that EVH1 domain is tethered to the membrane while the EVH2 domain is tethered to F-actin suggests that ENA/VASPmediated actin polymerization is likely influenced by mechanical forces. In addition, torque can possibly accumulate during actin polymerization if the relative rotation between the Ena/VASP tetramer and F-actin is restrained. Therefore, it is important to investigate how these mechanical constraints regulate Ena/VASP-mediated actin polymerization in future studies.
4.6. Advanced single-molecule experimental designs The implementation of various state-of-the-art single-molecule manipulation technologies has led to a better understanding of the effects of mechanical constraints on formin-mediated F-actin polymerization [27–29,31,32,40,44,45]. Most of the current technologies involve applying controlled, external tensile forces to formin-actin systems [27–29,31,32,40,44,45]. In the case of magnetic tweezers, the relative rotation between the formin and the F-actin can also be controlled [31,40]. However, the force and torque applied cannot be controlled simultaneously in any of these current technologies. Given the recent evidence that formin is also a torque sensor [31,40] new technologies need to be developed that can simultaneously apply well-controlled force and torque to F-actin and quantify the resulting formin-dependent actin polymerization under such constraints. While it is simpler to apply well-controlled, external force to a molecule, it is much more difficult to apply controlled, external torque. In this regard, a recent study implemented a modified magnetic tweezers design that can simultaneously apply force and torque to a molecule [58]. In this design, a super-paramagnetic bead is attached to the end of a DNA molecule and a nonmagnetic polystyrene bead is attached to the side of the DNA molecule. A rotating magnetic field with a constant angular velocity is applied, causing the rotation of the superparamagnetic bead and thus the rotation of the DNA molecule and the polystyrene bead with the same angular velocity. The force can be controlled by changing the distance between the magnets and the super-paramagnetic bead, while the torque can be controlled independently by changing the angular velocity of the rotating magnetic field. This design can be adapted into a transverse magnetic tweezers set up that can apply controlled force and torque during formin-mediated actin polymerization. In this setup the F-actin can be stretched in the imaging plane, as depicted in Fig. 3, which can be further integrated with fluorescence imaging of fluorescence-labeled F-actin with micropillar arrays. Finally, the experimental throughput can be increased by tracking multiple tethers in the same view area at the same time. Such integrated, multiplexing single-molecule manipulation and imaging technology has a great potential to reveal new insights into the molecular mechanisms underlying formin-dependent actin polymerization under mechanical constraints.
4.5. Mechanical regulation of F-actin destruction The mechanical regulation of formin-dependent F-actin polymerization has been well-studied; however, the mechanical principles governing F-actin destruction is not so well investigated [52–56]. The most common way for F-actin destruction is through depolymerization, where actin monomers dissociate from the F-actin ends. A recent study using single F-actin flow-stretching revealed that mDia1 remains associated at the barbed end of a depolymerizing F-actin, and the rate of depolymerization is significantly suppressed when the force applied to the F-actin is increased from 0 pN to 3 pN [28,32]. This observation suggests that force negatively regulates formin-mediated actin depolymerization. A second way of F-actin destruction is through F-actin filament cleavage, where an F-actin breaks into shorter fragments. Cofilin 6
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Fig. 3. Advanced single-molecule manipulation design for probing the torque and force effects on formin-mediated actin polymerization. (A) Illustrations of experimental designs to investigate force-dependent formin-mediated actin polymerization in a torque-free condition. In the sample channel, micro-pillar arrays (top left inset) are coupled with inactive myosin-coated non-magnetic polystyrene bead at pillar top. The pointed-end region of an actin filament is bound with the bead at pillar top. The barbed-end of the filament is bound with formin FH2 ring, while the FH1 domains are tethered to a superparamagnetic bead. A pair of magnets are used to generate forces along the filament, which magnetizes the bead in the direction perpendicular to the force direction and thus inhibits the rotation of the bead. A DNA linker (top panel) or a lipid bilayer (bottom panel) is added between the FH1 domains and the bead to enable the free rotation of formin FH2 ring around the actin filament during actin polymerization. When the FH1 domains are directly tethered to the superparamagnetic bead, rotation of formin around the actin filament can also be enabled using a cylindrically symmetric cone-shaped magnet (middle panel). In addition, the use of pillar array allows stretching of the filament in the focal plane, such that the elongating filament can be simultaneously visualized using fluorescence microscopy. (B) Illustration of an advanced magnetic tweezer design to control force and torque during formin-mediated actin polymerization. In the design, a pair of magnets is rotated with a constant angular velocity, which creates a rotating magnetic field that forces the side-attached super-paramagnetic bead and the filament to rotate with the same angular velocity. It further causes the rotation of a polystyrene bead attached to the FH1 domains of the formin dimer (bound to F-actin barbed end) with the same angular velocity. A DNA linker is used to make the polystyrene bead rotation unconstrained. The rotation drag to the polystyrene bead creates a torque at the formin-actin interface. The force can be controlled by changing the distance between the magnets and the superparamagnetic bead, while the torque can be controlled independently by changing the angular velocity of the rotating magnetic field. [5] A. Schonichen, M. Geyer, Fifteen formins for an actin filament: a molecular view on the regulation of human formins, Biochim. Biophys. Acta 1803 (2) (2010) 152–163. [6] J.E. Bear, F.B. Gertler, Ena/VASP: towards resolving a pointed controversy at the barbed end, J. Cell. Sci. 122 (Pt 12) (2009) 1947–1953. [7] D. Breitsprecher, A.K. Kiesewetter, J. Linkner, M. Vinzenz, T.E. Stradal, J.V. Small, U. Curth, R.B. Dickinson, J. Faix, Molecular mechanism of Ena/VASP-mediated actin-filament elongation, EMBO J. 30 (3) (2011) 456–467. [8] S.D. Hansen, R.D. Mullins, VASP is a processive actin polymerase that requires monomeric actin for barbed end association, J. Cell Biol. 191 (3) (2010) 571–584. [9] M. Krause, E.W. Dent, J.E. Bear, J.J. Loureiro, F.B. Gertler, Ena/VASP proteins: regulators of the actin cytoskeleton and cell migration, Annu. Rev. Cell Dev. Biol. 19 (2003) 541–564. [10] S. Romero, C. Le Clainche, D. Didry, C. Egile, D. Pantaloni, M.F. Carlier, Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis, Cell 119 (3) (2004) 419–429. [11] Y. Xu, J.B. Moseley, I. Sagot, F. Poy, D. Pellman, B.L. Goode, M.J. Eck, Crystal structures of a Formin Homology-2 domain reveal a tethered dimer architecture, Cell 116 (5) (2004) 711–723. [12] T. Otomo, D.R. Tomchick, C. Otomo, S.C. Panchal, M. Machius, M.K. Rosen, Structural basis of actin filament nucleation and processive capping by a formin homology 2 domain, Nature 433 (7025) (2005) 488–494. [13] D. Vavylonis, D.R. Kovar, B. O’Shaughnessy, T.D. Pollard, Model of formin-associated actin filament elongation, Mol. Cell 21 (4) (2006) 455–466. [14] P. Kursula, I. Kursula, M. Massimi, Y.H. Song, J. Downer, W.A. Stanley, W. Witke,
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