From the Cover: PNAS Plus: Cell-cycle regulation of formin-mediated actin cable assembly

Assembly of appropriately oriented actin cables nucleated by formin proteins is necessary for many biological processes in diverse eukaryotes. However, compared with knowledge of how nucleation of dendritic actin filament arrays by the actin-related protein-2/3 complex is regulated, the in vivo regulatory mechanisms for actin cable formation are less clear. To gain insights into mechanisms for regulating actin cable assembly, we reconstituted the assembly process in vitro by introducing microspheres functionalized with the C terminus of the budding yeast formin Bni1 into extracts prepared from yeast cells at different cell-cycle stages. EM studies showed that unbranched actin filament bundles were reconstituted successfully in the yeast extracts. Only extracts enriched in the mitotic cyclin Clb2 were competent for actin cable assembly, and cyclin-dependent kinase 1 activity was indispensible. Cyclin-dependent kinase 1 activity also was found to regulate cable assembly in vivo. Here we present evidence that formin cell-cycle regulation is conserved in vertebrates. The use of the cable-reconstitution system to test roles for the key actin-binding proteins tropomyosin, capping protein, and cofilin provided important insights into assembly regulation. Furthermore, using mass spectrometry, we identified components of the actin cables formed in yeast extracts, providing the basis for comprehensive understanding of cable assembly and regulation.Eukaryotic cells contain populations of actin structures with distinct architectures and protein compositions, which mediate varied cellular processes (1). Understanding how F-actin polymerization is regulated in time and space is critical to understanding how actin structures provide mechanical forces for corresponding biological processes. Branched actin filament arrays, which concentrate at sites of clathrin-mediated endocytosis (2, 3) and at the leading edge of motile cells (4), are nucleated by the actin-related protein-2/3 (Arp2/3) complex. In contrast, bundles of unbranched actin filaments, which sometimes mediate vesicle trafficking or form myosin-containing contractile bundles, often are nucleated by formin proteins (5–14).Much has been learned about how branched actin filaments are polymerized by the Arp2/3 complex and how these filaments function in processes such as endocytosis (2, 15). In contrast, relatively little is known about how actin cables are assembled under physiological conditions. In previous studies, branched actin filaments derived from the Arp2/3 complex have been reconstituted using purified proteins (16–19) or cellular extracts (20–25). When microbeads were coated with nucleation-promoting factors for the Arp2/3 complex and then were incubated in cell extracts, actin comet tails were formed by sequential actin nucleation, symmetry breaking, and tail elongation. Importantly, the motility behavior of F-actin assembled by the Arp2/3 complex using defined, purified proteins differs from that of F-actin assembled by the Arp2/3 complex in the full complexity of cytoplasmic extracts (19, 26–28).Formin-based actin filament assembly using purified proteins also has been reported (29, 30). However, reconstitution of formin-derived actin cables under the more physiological conditions represented by cell extracts has not yet been reported.The actin nucleation activity of formin proteins is regulated by an inhibitory interaction between the N- and C-terminal domains, which can be released when GTP-bound Rho protein binds to the formin N-terminal domain, allowing access of the C terminus (FH1-COOH) to actin filament barbed ends (31–40). In yeast, the formin Bni1 N terminus also has an inhibitory effect on actin nucleation through binding to the C terminus (41).Interestingly, several recent reports provided evidence for cell-cycle regulation of F-actin dynamics in oocytes and early embryos (42–45). However, which specific types of actin structures are regulated by the cell cycle and what kind of nucleation factors and actin interacting-proteins are involved remain to be determined.Here, we report a reconstitution of actin cables in yeast extracts from microbeads derivatized with Bni1 FH1-COOH, identifying the proteins involved, increasing the inventory of the proteins that regulate actin cable dynamics and establishing that the actin cable reconstitution in cytoplasmic extracts is cell-cycle regulated.     

Mechanical stimulation induces formin-dependent assembly of a perinuclear actin rim

Cells constantly sense and respond to mechanical signals by reorganizing their actin cytoskeleton. Although a number of studies have explored the effects of mechanical stimuli on actin dynamics, the immediate response of actin after force application has not been studied. We designed a method to monitor the spatiotemporal reorganization of actin after cell stimulation by local force application. We found that force could induce transient actin accumulation in the perinuclear region within ∼2 min. This actin reorganization was triggered by an intracellular Ca²⁺ burst induced by force application. Treatment with the calcium ionophore A23187 recapitulated the force-induced perinuclear actin remodeling. Blocking of actin polymerization abolished this process. Overexpression of Klarsicht, ANC-1, Syne Homology (KASH) domain to displace nesprins from the nuclear envelope did not abolish Ca²⁺-dependent perinuclear actin assembly. However, the endoplasmic reticulum- and nuclear membrane-associated inverted formin-2 (INF2), a potent actin polymerization activator (mutations of which are associated with several genetic diseases), was found to be important for perinuclear actin assembly. The perinuclear actin rim structure colocalized with INF2 on stimulation, and INF2 depletion resulted in attenuation of the rim formation. Our study suggests that cells can respond rapidly to external force by remodeling perinuclear actin in a unique Ca²⁺- and INF2-dependent manner.Cells can sense and adapt to their physical microenvironment through specific mechanosensing mechanisms. These properties are often mediated by the actin cytoskeleton, which can be modulated by a wide range of forces. Fluid shear stress, for example, induces actin stress fiber assembly and realignment along the direction of flow (1–4), whereas the cyclic stretch of an elastic substrate induces a reorientation of stress fibers under some angle to the direction of stretch (5–8). Applying mechanical force to cells by a microneedle results in focal adhesion growth and activation of formin-type actin nucleators (9, 10). Similarly, local application of force through fibronectin or collagen-coated beads trapped by optical or magnetic tweezers leads to the local reorganization of the actin cytoskeleton. This response is associated with reinforcement of bead attachment (11), recruitment of additional actin-associated proteins (12), and activation of a variety of signaling pathways (13–17). Most studies to date have explored the effects of force on actin structures directly associated with the sites of force application, such as focal adhesions and stress fibers. However, the immediate effect of force on the assembly of actin structures distal from the sites of force application has not been assessed. Such process is despite distal effects having potential implications in the transduction of local forces from the cell periphery to nuclear events (18).In this study, we used a local mechanical force application device and examined the large-scale actin reorganization during and after force application. Remarkably, we identified reversible actin polymerization in the perinuclear region within 1 min after mechanical stimulation. Intracellular Ca²⁺ bursts were found to be essential for the perinuclear actin response. Furthermore, we showed that a potent actin polymerization factor, inverted formin-2 (INF2), was involved in the perinuclear actin remodeling. Specifically, INF2 colocalized with a transient actin structure in the perinuclear region. A reduction in the level of INF2 resulted in the attenuation of this actin remodeling process. This work reveals a previously unidentified mechanotransduction response, whereby external mechanical stimulation induces a rapid transient perinuclear actin polymerization mediated by Ca²⁺ and formin.     

Structure of the formin-interaction domain of the actin nucleation-promoting factor Bud6

Formin proteins and their associated factors cooperate to assemble unbranched actin filaments in diverse cellular structures. The Saccharomyces cerevisiae formin Bni1 and its associated nucleation-promoting factor (NPF) Bud6 generate actin cables and mediate polarized cell growth. Bud6 binds to both the tail of the formin and G-actin, thereby recruiting monomeric actin to the formin to create a nucleation seed. Here, we structurally and functionally dissect the nucleation-promoting C-terminal region of Bud6 into a Bni1-binding “core” domain and a G-actin binding “flank” domain. The ∼2-Å resolution crystal structure of the Bud6 core domain reveals an elongated dimeric rod with a unique fold resembling a triple-helical coiled-coil. Binding and actin-assembly assays show that conserved residues on the surface of this domain mediate binding to Bni1 and are required for NPF activity. We find that the Bni1 dimer binds two Bud6 dimers and that the Bud6 flank binds a single G-actin molecule. These findings suggest a model in which a Bni1/Bud6 complex with a 2:4 subunit stoichiometry assembles a nucleation seed with Bud6 coordinating up to four actin subunits.The assembly of diverse filamentous actin arrays in cells is dependent on machinery that catalyzes the otherwise inefficient step of actin nucleation. A variety of actin nucleators and nucleation-promoting factors (NPFs) have now been identified, including Arp2/3 complex, WASp/WAVE family members, formins, Spire, Cobl, Lmod, JMY, adenomatous polyposis coli (APC), and Bud6 (reviewed in refs. 1–4). Although each is unique in its detailed mechanism of actin assembly, many of these factors are surprisingly related in their overall strategy for promoting polymerization from a pool of free actin monomers. Additionally, efficient actin assembly often requires cooperation between an actin nucleator and one or more NPFs. A majority of these nucleators or NPFs contain multiple WASP homology-2 (WH2) domains, a short (17–27 aa) motif that binds actin monomers (2). The need for nucleators to recruit multiple actin monomers stems from the fact that actin dimers and trimers are extremely unstable, short-lived species. The smallest stable actin species is a tetramer, which has a K_d of 0.14 μM (5). The classic Arp2/3 complex exploits interactions with WH2-containing WASP-family NPFs in its mechanism of actin filament nucleation. Together, the Arp2 and Arp3 subunits in the complex resemble a short-pitched actin dimer and bind to two NPF molecules (6–8), each of which brings in at least one actin monomer. This is thought to generate a four-actin cross-filament seed for a daughter filament that rapidly polymerizes at an angle of 70° to the mother filament (9). Other WH2-containing proteins can act independently to nucleate actin filaments, likely by arraying multiple actin subunits into a nucleation seed. For example, Cobl has three WH2 domains and an unusually long linker sequence separating its second and third WH2 domains, which is critical for nucleation (10). This has led to the proposal that Cobl arranges monomers into a cross-filament trimer that serves as a seed for polymerization.Formin-family nucleators generally lack WH2 domains, and they also differ from the Arp2/3 complex in that they nucleate unbranched actin filaments, such as those found in cytokinetic rings, filopodia, stress fibers, and yeast actin cables (4, 11). Formins vary in their domain structure, reflecting their diverse cellular roles and mechanisms of regulation, but all contain the highly conserved formin homology-2 (FH2) domain. The FH2 domain consists of two rod-shaped subdomains tied together in a head-to-tail arrangement by flexible linkers to form a closed ring. Each side of the dimer contains two actin-binding surfaces, allowing the FH2 dimer to organize two or three actin subunits into a filament-like orientation that can function as a nucleus for filament polymerization (12, 13). Formins associate with the “barbed” end of a nascent filament, and the flexible nature of the FH2 dimer affords it the surprising ability to “stair-step” on the elongating barbed end as new actin subunits are incorporated (4, 14). Elongation is also facilitated by the formin homology-1 (FH1) domain, a segment immediately adjacent to the FH2 domain that contains multiple proline-rich motifs and can accelerate elongation by recruiting profilin-bound actin subunits to the site of incorporation at the barbed end of the growing filament (15, 16).Formins were initially proposed to nucleate actin filaments solely using their FH2 domains, perhaps by stabilizing transiently formed dimers or trimers (17) because the isolated FH2 domain lacks significant affinity for actin monomers (12). However, recent work has revealed that an adjacent C-terminal tail region, often containing the diaphanous autoregulatory domain (DAD), directly binds actin monomers and works together with the FH2 domain to stimulate nucleation (18). Other formins may use a WH2-like element distinct from the DAD domain for this purpose (19, 20). Furthermore, a growing number of formins have been shown to bind directly to actin monomer-recruiting NPFs. For example, the Drosophila formin Cappuccino and its mammalian counterparts Fmn1 and Fmn2 bind to Spire, an actin nucleator that contains an array of four WH2 domains (21). The mammalian formin mDia1 interacts with APC protein, an actin nucleator that binds monomers but does not have identifiable WH2 domains (22). In budding yeast, the formin Bni1 binds Bud6, which also binds actin monomers but does not have a clearly recognizable WH2 domain (23).Bud6 localizes to the bud tip and neck, and it was first identified in a yeast two-hybrid screen for actin-interacting proteins (24). In Schizosaccharomyces pombe, the microtubule plus end-associated protein Tea1, formin For3, and Bud6 form a large polarity complex that resides at the cell tips and promotes localized actin cable assembly, and the triple knockout of these three genes leads to severe defects in cell polarity (25). Bud6 also contributes to the maintenance of septin-dependent diffusion barriers in the endoplasmic reticulum and nuclear membranes, which limit membrane protein diffusion between the mother and daughter cell compartments (26, 27). The N-terminal half of Bud6 is required for its in vivo localization and for its function in cortical capture of astral microtubule ends (28, 29). It has also been shown to bind microtubules directly (28, 29). The C-terminal half (residues 489–788) directly facilitates actin filament assembly by the formin Bni1 (23). Bud6 enhances the nucleation phase, rather than the elongation phase, of Bni1-mediated actin filament assembly, and this NPF effect requires separable interactions of Bud6 with Bni1 and with actin monomers (30).To understand better how Bud6 functions together with Bni1 in actin assembly, we dissected the structural and functional properties of a C-terminal fragment of Bud6 that contains NPF activity (c-Bud6, residues 550–788). We find that c-Bud6 can functionally be divided into two parts: a trypsin stable core (residues 550–688) that contains the Bni1 binding site and a flank (residues 699–788) that binds to actin monomers. Although the core domain retains the ability to bind Bni1, it inhibits, rather than stimulates, actin nucleation by Bni1, likely because it obstructs the actin monomer recruitment activity of the Bni1 tail region. The crystal structure of the trypsin stable Bud6 core reveals a unique rod-shaped dimeric fold. Conserved surfaces at either end of the core domain and at its center are critical for Bni1 binding and NPF activity. Through a series of native gel shift, size exclusion chromatography (SEC) multiangle light scattering (MALS), and isothermal titration calorimetry (ITC) experiments, we determined the stoichiometry of association of Bud6 with Bni1 and with actin monomers. These structural and functional data inform an emerging model for the mechanism of actin nucleation by this nucleator/NPF pair.     

Orientation-specific responses to sustained uniaxial stretching in focal adhesion growth and turnover

Cells are mechanosensitive to extracellular matrix (ECM) deformation, which can be caused by muscle contraction or changes in hydrostatic pressure. Focal adhesions (FAs) mediate the linkage between the cell and the ECM and initiate mechanically stimulated signaling events. We developed a stretching apparatus in which cells grown on fibronectin-coated elastic substrates can be stretched and imaged live to study how FAs dynamically respond to ECM deformation. Human bone osteosarcoma epithelial cell line U2OS was transfected with GFP-paxillin as an FA marker and subjected to sustained uniaxial stretching. Two responses at different timescales were observed: rapid FA growth within seconds after stretching, and delayed FA disassembly and loss of cell polarity that occurred over tens of minutes. Rapid FA growth occurred in all cells; however, delayed responses to stretch occurred in an orientation-specific manner, specifically in cells with their long axes perpendicular to the stretching direction, but not in cells with their long axes parallel to stretch. Pharmacological treatments demonstrated that FA kinase (FAK) promotes but Src inhibits rapid FA growth, whereas FAK, Src, and calpain 2 all contribute to delayed FA disassembly and loss of polarity in cells perpendicular to stretching. Immunostaining for phospho-FAK after stretching revealed that FAK activation was maximal at 5 s after stretching, specifically in FAs oriented perpendicular to stretch. We hypothesize that orientation-specific activation of strain/stress-sensitive proteins in FAs upstream to FAK and Src promote orientation-specific responses in FA growth and disassembly that mediate polarity rearrangement in response to sustained stretch.Tissues of the pulmonary, circulatory, musculoskeletal, digestive, and renal systems are subject to mechanical perturbations such as cyclic or continuous stretch. Cells within these tissues are mechanosensitive, responding to these mechanical inputs by changing ion channel configurations (1–3), cytoskeleton organization (4–6), mRNA splicing (7), gene expression (8–10), and posttranslational protein modification (11–13). Although many forms of mechanical stimuli occur physiologically, the most well studied are the responses of cells to cyclic stretch to specifically simulate the contraction/relaxation cycles that occur within the cardiovascular and pulmonary systems (14–17). However, sustained stretch also commonly occurs in tissues—e.g., when injury-induced swelling causes local hydrostatic pressure increase (18), in long-term blood pressure increase (16, 19), during prolonged muscle contraction (20, 21), or when the bladder retains large volume of urine (22, 23). In such situations, tissue stretching may last for minutes or hours without regular relaxation intervals, and likely generates cellular responses that lead to tissue adaptation. However, the cellular responses to sustained stretching are not well studied.Cells exhibit stereotypical morphological responses to stretch. Cyclic uniaxial stretch induces cells to reorient their long axes perpendicular to the direction of stretch. This process is accompanied by a similar reorientation of actomyosin stress fibers and integrin-mediated focal adhesions (FAs) and is thought to minimize tissue resistance to stretch (5, 6, 24, 25). FAs serve as mechanical conduits, transmitting forces generated by stress fibers to the ECM to drive cell movement or ECM remodeling, and also transmitting forces generated in tissues into the cell. The mechanosensitivity of FA assembly and downstream signaling is well documented, and thus they are prime candidates for mediating cellular responses to stretch (26). Indeed, the cytoskeletal reorientation that occurs in response to cyclic stretch requires several proteins that localize to FA, including integrins (27), zyxin (28), paxillin (25), Src, and p130 Crk-associated substrate (p130Cas) (29), and involves sliding reorganization of FAs (6). However, the effects of sustained stretch on FA organization and dynamics have not been explored.Here, we sought to characterize the dynamic response of FAs and cell morphologies to sustained uniaxial stretch. We found FAs to be directionally sensitive to sustained stretching. FA disassembly and loss of cell polarity occurred in cells with their long axes perpendicular to stretch, but not in cells with their long axes parallel to stretch. We show that FA kinase (FAK), Src family kinases (SFKs), and calpain 2 all contribute to FA disassembly and loss of cell polarity, and that FAK is activated specifically in FAs oriented perpendicular to stretch. We hypothesize that putative stress/strain sensing proteins in FAs upstream to FAK and Src align with the long axes of FAs, whereas a majority of FAs aligns with the cellular long axis, resulting in orientation-specific responses. Our findings show that sensitivity of strain/stress orientation at the molecular level propagates up to regulate cell polarity in response to sustained mechanical stimulus.     

Actomyosin dynamics drive local membrane component organization in an in vitro active composite layer

The surface of a living cell provides a platform for receptor signaling, protein sorting, transport, and endocytosis, whose regulation requires the local control of membrane organization. Previous work has revealed a role for dynamic actomyosin in membrane protein and lipid organization, suggesting that the cell surface behaves as an active composite composed of a fluid bilayer and a thin film of active actomyosin. We reconstitute an analogous system in vitro that consists of a fluid lipid bilayer coupled via membrane-associated actin-binding proteins to dynamic actin filaments and myosin motors. Upon complete consumption of ATP, this system settles into distinct phases of actin organization, namely bundled filaments, linked apolar asters, and a lattice of polar asters. These depend on actin concentration, filament length, and actin/myosin ratio. During formation of the polar aster phase, advection of the self-organizing actomyosin network drives transient clustering of actin-associated membrane components. Regeneration of ATP supports a constitutively remodeling actomyosin state, which in turn drives active fluctuations of coupled membrane components, resembling those observed at the cell surface. In a multicomponent membrane bilayer, this remodeling actomyosin layer contributes to changes in the extent and dynamics of phase-segregating domains. These results show how local membrane composition can be driven by active processes arising from actomyosin, highlighting the fundamental basis of the active composite model of the cell surface, and indicate its relevance to the study of membrane organization.The cell surface mediates interactions between the cell and the outside world by serving as the site for signal transduction. It also facilitates the uptake and release of cargo and supports adhesion to substrates. These diverse roles require that the cell surface components involved in each function are spatially and temporally organized into domains spanning a few nanometers (nanoclusters) to several micrometers (microdomains). The cell surface itself may be considered as a fluid–lipid bilayer wherein proteins are embedded (1). In the living cell, this multicomponent system is supported by an actin cortex, composed of a branched network of actin and a collection of filaments (2–4).Current models of membrane organization fall into three categories: those invoking lipid–lipid and lipid–protein interactions in the plasma membrane [e.g., the fluid mosaic model (1, 5) and the lipid raft hypothesis (6)], or those that appeal to the membrane-associated actin cortex (e.g., the picket fence model) (7), or a combination of these (8, 9). Although these models based on thermodynamic equilibrium principles have successfully explained the organization and dynamics of a range of membrane components and molecules, there is a growing class of phenomena that appears inconsistent with chemical and thermal equilibrium, which might warrant a different explanation. These include aspects of the organization and dynamics of outer leaflet glycosyl-phosphatidylinositol-anchored proteins (GPI-anchored proteins) (10–13), inner leaflet Ras proteins (14), and actin-binding transmembrane proteins (13, 15, 16).Recent experimental and theoretical work has shown that these features can be explained by taking into account that many cortical and membrane proteins are driven by ATP-consuming processes that drive the system out of equilibrium (13, 15, 17). The membrane models mentioned above have by-and-large neglected this active nature of the actin cortex where actin filaments are being continuously polymerized and depolymerized (18–21), in addition to being persistently acted upon by a variety of myosin motors (22–24) that consume ATP and exert contractile stresses on cortical actin filaments, continually remodeling the architecture of the cortex (4, 21, 25). These active processes in turn can generate tangential stresses and currents on the cell surface, which could drive the dynamics and local composition of membrane components at different scales (22, 26–29).Actin polymerization is proposed to be driven at the membrane by two nucleators, the Arp2/3 complex, which creates a densely branched network, as well as formins that nucleate filaments (18, 21, 30). A number of myosin motors are also associated with the juxtamembranous actin cortex, of which nonmuscle myosin II is the major component in remodeling the cortex and creating actin flows (4, 23, 25, 26, 31, 32). Based on our observations that the clustering of cell surface components that couple directly or indirectly to cortical actin [e.g., GPI-anchored proteins, proteins of the Ezrin, Radaxin, or Moesin (ERM) family (13, 15)] depends on myosin activity, we proposed that this clustering arises from the coupling to contractile actomyosin platforms (called “actin asters”) produced at the cortex (15, 33).A coarse-grained theory describing this idea has been put forward and corroborated by the verification of its key predictions in live cells (15, 33), but a systematic identification of the underlying microscopic processes is lacking. Given the complexity of numerous processes acting at the membrane of a living cell, we use an in vitro approach to study the effect of an energy-consuming actomyosin network on the dynamics of membrane molecules that directly interact with filamentous actin.A series of in vitro studies have explored the organization of confined, dynamic filaments (both actin and microtubules) (34–39) or the role of actin architecture on membrane organization (40–46). Indeed, these studies have yielded insights into the nontrivial emergent configurations that mixtures of polar filaments and motors can adopt when fueled by ATP (34–37), in particular constitutively remodeling steady states that display characteristics of active mechanics (38, 39, 47). However, the effect of linking these mechanics to the confining lipid bilayer and its organization has not been studied.The consequences of actin polymerization on membrane organization, in particular on giant unilamellar vesicles (GUVs), have been addressed in a number of studies on the propulsion of GUVs by an actin comet tail (40, 45, 46). In those experiments, the apparent advection of membrane bound ActA or WASP toward the site of actin polymerization is mainly due to the change in binding affinity of WASP to actin through Arp2/3 (44) and the spherical geometry resulting in the drag of actin to one pole of the vesicle after symmetry break of the actin shell. That this dynamic process changes the bulk properties of the bilayer, namely the critical temperature of a phase-separating lipid bilayer, was shown by Liu and Fletcher (40) when the actin nucleator N-WASP was connected to a lipid species (PIP₂) that was capable of partitioning into one of the two phases.Besides these pioneering studies on the effects of active processes on membrane organization, little was done to directly test the effect of active lateral stresses as well as actomyosin remodeling at the membrane, particularly on the dynamics and organization of membrane-associated components.To this end, we build an active composite in vitro by stepwise addition of components: a supported lipid bilayer with an actin-binding component, actin filaments, and myosin motors. By systematically varying the concentrations of actin and myosin as well as the average actin filament length, we find distinct states of actomyosin organization at the membrane surface upon complete ATP consumption. More importantly, we find that the ATP-fueled contractile actomyosin currents induce the transient accumulation of actin-binding membrane components. As predicted, the active mechanics of actin and myosin at physiologically relevant ATP concentrations drives the system into a nonequilibrium steady state with anomalous density fluctuations and the transient clustering of actin-binding components of the lipid bilayer (15, 33). Finally, connection of this active layer of actomyosin to a phase-segregating bilayer, influences its phase behavior and coarsening dynamics.     

Chronophin coordinates cell leading edge dynamics by controlling active cofilin levels

Cofilin, a critical player of actin dynamics, is spatially and temporally regulated to control the direction and force of membrane extension required for cell locomotion. In carcinoma cells, although the signaling pathways regulating cofilin activity to control cell direction have been established, the molecular machinery required to generate the force of the protrusion remains unclear. We show that the cofilin phosphatase chronophin (CIN) spatiotemporally regulates cofilin activity at the cell edge to generate persistent membrane extension. We show that CIN translocates to the leading edge in a PI3-kinase–, Rac1-, and cofilin-dependent manner after EGF stimulation to activate cofilin, promotes actin free barbed end formation, accelerates actin turnover, and enhances membrane protrusion. In addition, we establish that CIN is crucial for the balance of protrusion/retraction events during cell migration. Thus, CIN coordinates the leading edge dynamics by controlling active cofilin levels to promote MTLn3 cell protrusion.Cofilin is one crucial mediator of actin cytoskeletal dynamics during cell motility (1–5). At the cell edge, cofilin severs F-actin filaments, generating substrates for Arp2/3-mediated branching activity and contributing to F-actin depolymerization by creating a new pointed end and F-actin assembly by increasing the pool of polymerization-competent actin monomers (G-actin) (6, 7). Because of its ability to sever actin filaments and thus, modulate actin dynamics, the precise spatial and temporal regulation of cofilin activity at the cell leading edge is crucial to cell protrusion, chemotaxis, and motility both in vitro and in vivo (2, 8–13). Misregulation of cofilin activity and/or expression is directly related to diseases, including tumor metastasis (14–18) and Alzheimer’s disease (19).Several mechanisms regulate tightly the activation of cofilin in response to upstream stimuli, including interaction with phosphatidylinositol (4,5)-bisphosphate (20–22), local pH changes (23, 24), and phosphorylation at a single regulatory serine (Ser3) (8, 25). The phosphorylation of cofilin, leading to its inactivation, is catalyzed by two kinase families: the LIM-kinases [LIMKs(Lin11, Isl-1, and Mec-3 domain)] and the testicular kinases (25–27). Two primary families of ser/thr phosphatases dephosphorylate and reactivate the actin-depolymerizing and -severing functions of cofilin: slingshot (SSH) (28) and chronophin (CIN) (29).SSH was identified as a cofilin phosphatase through genetic studies in Drosophila (28). The most active and abundant SSH isoform, SSH-1L, has been implicated in such biological processes as cell division, growth cone motility/morphology, neurite extension, and actin dynamics during membrane protrusion (30). SSH dephosphorylates a number of actin regulatory proteins in addition to cofilin, including LIMK1 (31) and Coronin 1B (32). CIN is a haloacid dehydrogenase-type phosphatase, a family of enzymes with activity in mammalian cells that has been poorly characterized. CIN dephosphorylates a very limited number of substrates (33) and as opposed to SSH, has little phosphatase activity toward LIMK both in vitro and in vivo; thus, it seems to be the more specific activator of cofilin (29, 30). CIN exhibits several predicted interaction motifs potentially linking it to regulation by PI3-kinase and phospholipase Cγ (PLCγ), both of which have been implicated in signaling to cofilin activation in vivo in MTLn3 adenocarcinoma cells (10, 34). CIN has been involved in cell division (29), cofilin–actin rod formation in neurons (35), and chemotaxing leukocytes (36, 37). The molecular mechanisms that control the activity and localization of CIN in cells are still not well-understood. In neutrophils, CIN mediates cofilin dephosphorylation downstream of Rac2 (36), and stimulation of protease-activated receptor2 results in recruitment of CIN and cofilin at the cell edge by β-arrestins to promote localized generation of free actin barbed ends, membrane protrusion, and chemotaxis (37). Chemotaxis to EGF by breast tumor cells is directly correlated with cancer cell invasion and metastasis (38, 39). Although cofilin activity is required for tumor cell migration, the contribution(s) of CIN to the regulation of actin dynamics at the leading edge has not yet been investigated.The importance of cofilin in regulating tumor cell motility has been extensively studied using MTLn3 mammary carcinoma cells as a model system. The initial step of MTLn3 cell chemotaxis to EGF consists of a biphasic actin polymerization response resulting from two peaks of free actin barbed end formation (34, 40, 41). The first or early peak of actin polymerization occurs at 1 min after EGF stimulation and requires both cofilin and PLCγ activities (34), but it is not dependent on cofilin dephosphorylation (42). This first transient allows the cells to sense EGF gradients and initiate small-membrane protrusions (11). The second or late peak of actin polymerization occurs at 3 min and is dependent on both cofilin and PI3-kinase activities (43, 44). Cofilin activity in this late transient has been associated with full protrusion of lamellipodia (34). The mechanism by which cofilin becomes activated at the 3-min peak has not been identified, although it is likely to involve the phosphoregulation of Ser3 (42, 45).In this work, we determine the molecular mechanisms involved in the full protrusion of the leading edge upon EGF stimulation. We have identified CIN as a critical regulator of cofilin activation to coordinate leading edge dynamics. Our results yield insights into how CIN controls cell protrusion, a key step in the process of cell migration and metastasis.     

Talin determines the nanoscale architecture of focal adhesions

Insight into how molecular machines perform their biological functions depends on knowledge of the spatial organization of the components, their connectivity, geometry, and organizational hierarchy. However, these parameters are difficult to determine in multicomponent assemblies such as integrin-based focal adhesions (FAs). We have previously applied 3D superresolution fluorescence microscopy to probe the spatial organization of major FA components, observing a nanoscale stratification of proteins between integrins and the actin cytoskeleton. Here we combine superresolution imaging techniques with a protein engineering approach to investigate how such nanoscale architecture arises. We demonstrate that talin plays a key structural role in regulating the nanoscale architecture of FAs, akin to a molecular ruler. Talin diagonally spans the FA core, with its N terminus at the membrane and C terminus demarcating the FA/stress fiber interface. In contrast, vinculin is found to be dispensable for specification of FA nanoscale architecture. Recombinant analogs of talin with modified lengths recapitulated its polarized orientation but altered the FA/stress fiber interface in a linear manner, consistent with its modular structure, and implicating the integrin–talin–actin complex as the primary mechanical linkage in FAs. Talin was found to be ∼97 nm in length and oriented at ∼15° relative to the plasma membrane. Our results identify talin as the primary determinant of FA nanoscale organization and suggest how multiple cellular forces may be integrated at adhesion sites.Cell adhesion to the ECM is a highly coordinated process that involves ECM-specific recognition by integrin transmembrane receptors, and their aggregation with numerous cytoplasmic proteins into dense supramolecular complexes called focal adhesions (FAs) (1). Actin stress fibers terminate at FAs where actomyosin contractility is transmitted to the ECM, generating traction (2–5). Mechanical tension impinging on each FA is implicated in key steps including the elongation, reinforcement, and maintenance of the FA structures (6). FA mechanotransduction is a major aspect of cellular microenvironment sensing with wide-ranging consequences in physiological and pathological processes (7–10). However, molecular-scale spatial parameters that specify FA nanoscale organization have been difficult to access experimentally. Nevertheless, these are essential to understand how mechanosensitivity arises within such complex molecular machines (11–15).Previously 3D superresolution fluorescence microscopy has unveiled the nanoscale organization of major FA components, whereby a core region of ∼30 nm interposes between the integrin and the actin cytoskeleton along the vertical (z) axis (16). The FA core consists of a membrane-proximal layer that contains signaling proteins such as FAK (focal adhesion kinase) and paxillin, an intermediate zone that contains force-transduction proteins such as talin and vinculin, and a stress fiber interfacial zone that contains actin-associated proteins such as VASP (vasodilator-stimulated protein) and α-actinin. Although such multilaminar architecture signifies a certain degree of compartmentalization within FAs that may serve to spatially constrain protein–protein interactions and dynamics, the structural connectivity, the molecular configuration and geometry of FA proteins, and the molecular basis of their higher-order organization remain unclear.Proteomic and interactome analysis of the integrin adhesome have uncovered several direct and multitier connections between integrins and actin (17–20). This suggests that multiple highly interconnected protein–protein interactions could collectively self-organize into FA structures; such redundancy could also account for the remarkable mechanical robustness of FAs after cellular disruption or perturbation (21). Alternatively, a specific FA component may play a dominant role in regulating FA architecture. Aspects of both scenarios may also act cooperatively or function at distinct stages of FA assembly and maturation. Superresolution microscopy of cells expressing fluorescent protein (FP)-tagged FA components has revealed that talin, a large cytoskeletal adaptor protein, adopts a highly polarized orientation in FAs (16), with the N terminus residing in the membrane-proximal layer and the C terminus elevated by z ∼30 nm to the FA/stress fiber interfacial zone. This led us to hypothesize that an array of integrin–talin–actin linkages may vertically span the FA core, serving a structural role in determining FA architecture (16).To test this hypothesis, we sought to perturb FAs by substituting endogenous talin with recombinant analogs having modified lengths. These were generated by retaining both the N-terminal FERM (band 4.1/Ezrin/Radixin/Moesin) and C-terminal THATCH (Talin/HIP1R/Sla2p Actin-tethering C-terminal Homology, or R13) domains but with selective deletion of the multiple helical bundles within the central region of talin. By using a siRNA-mediated knockdown/rescue approach, we found that such talin analogs were able to support FA formation, clustering of activated integrins, and linkages to the actin cytoskeleton. By mapping the z-position of the FPs tagged at either the N or the C termini, we show that talin and its analogs are linearly extended and oriented in FAs, with their lengths regulating FA nanoscale organization. Chimeric-talin analogs with a 30-nm spacer insertion are also able to support FA assembly, facilitating the precise determination of talin geometry in FAs. Our results indicate that talin is oriented at 15° relative to the plasma membrane, measuring ∼97 nm end to end. FA nanoscale architecture in vinculin-null mouse embryonic fibroblasts (MEFs) retained its stratified organization and talin polarization similar to that in other cell types, suggesting that vinculin is dispensable for the specification of FA architecture. Our measurements demonstrate how the integrin–talin–actin module serves as the primary, and surprisingly modular, structural and tension-bearing core of FAs and geometrically define how such complexes could integrate multiple cellular forces at adhesion sites.     

Tension modulates actin filament polymerization mediated by formin and profilin

Formins promote processive elongation of actin filaments for cytokinetic contractile rings and other cellular structures. In vivo, these structures are exposed to tension, but the effect of tension on these processes was unknown. Here we used single-molecule imaging to investigate the effects of tension on actin polymerization mediated by yeast formin Bni1p. Small forces on the filaments dramatically slowed formin-mediated polymerization in the absence of profilin, but resulted in faster polymerization in the presence of profilin. We propose that force shifts the conformational equilibrium of the end of a filament associated with formin homology 2 domains toward the closed state that precludes polymerization, but that profilin–actin associated with formin homology 1 domains reverses this effect. Thus, physical forces strongly influence actin assembly by formin Bni1p.A host of proteins regulate the actin cytoskeleton by controlling filament nucleation, elongation, capping, branching, and bundling. Members of the formin family of proteins nucleate new filaments and remain processively attached to barbed ends while promoting the elongation of unbranched filaments (1, 2). Formin homology (FH)2 domains form a donut-shaped head-to-tail homodimer that encircles the fast-growing barbed end of actin filaments and promotes nucleation and polymerization (1, 3). When an actin monomer binds to the barbed end of a filament, one FH2 domain steps onto the new subunit, allowing the formin to remain attached to the filament through thousands of cycles of subunit addition (Fig. 1A) (2, 4). The FH2-bound end of the filament binds incoming actin monomers when in an “open” conformation but not in the “closed” conformation. As a consequence, FH2 domains slow barbed-end elongation compared with free barbed ends, a phenomenon termed “gating” (1, 5). Despite gating, FH2 domains can promote rapid filament elongation when coupled to FH1 domains (6), which are located N-terminal to the FH2 domain (7). Multiple polyproline tracks in FH1 domains bind the small actin-binding protein profilin, which mediates association of several profilin–actin complexes in close proximity to the end of a filament. Diffusive motions of the FH1 domain transfer actin rapidly to the filament barbed end (5), allowing elongation at rates faster than addition of subunits from the bulk phase.Open in a separate windowFig. 1.Formin-mediated actin filament polymerization in actin curtains. (A) Schematic of the reaction pathways for actin filament elongation by formin. FH2 dimers are shown in red for the closed conformation and green for the open conformation. End-on views of the filament illustrate the hypothesis that the closed conformation corresponds to a 180° pitch of the filament, whereas the open conformation has a 167° pitch (25). (B) Schematic of actin curtains. Biotinylated formin (Bni1p) is anchored via streptavidin to a lipid bilayer and polymerizes actin filaments that are aligned along nanofabricated barriers by solvent flow, allowing visualization by total internal reflection fluorescence microscopy.Actin filaments are subject to tension in cells, yet the influence of tension on formin-mediated polymerization was unknown, and theories predicted different outcomes in the absence and presence of profilin. Kozlov and colleagues proposed that the elasticity of the FH2 domain and the formin–barbed end binding energy govern the polymerization rate (8, 9). Their simulations suggested that tension increases the rate of polymerization by lowering the activation barrier for subunit addition and energetically favoring FH2 domain stepping onto the incoming subunit (8). Vavylonis et al. (10) postulated that force-induced stretching of FH1 domains slows the transfer of profilin–actin to the growing filament, so tension would inhibit polymerization in the presence of profilin. Here we describe experiments to test these predictions. We find that the effects of force are opposite both predictions for budding yeast formin Bni1p.     

Site-specific cation release drives actin filament severing by vertebrate cofilin

Actin polymerization powers the directed motility of eukaryotic cells. Sustained motility requires rapid filament turnover and subunit recycling. The essential regulatory protein cofilin accelerates network remodeling by severing actin filaments and increasing the concentration of ends available for elongation and subunit exchange. Although cofilin effects on actin filament assembly dynamics have been extensively studied, the molecular mechanism of cofilin-induced filament severing is not understood. Here we demonstrate that actin filament severing by vertebrate cofilin is driven by the linked dissociation of a single cation that controls filament structure and mechanical properties. Vertebrate cofilin only weakly severs Saccharomyces cerevisiae actin filaments lacking this “stiffness cation” unless a stiffness cation-binding site is engineered into the actin molecule. Moreover, vertebrate cofilin rescues the viability of a S. cerevisiae cofilin deletion mutant only when the stiffness cation site is simultaneously introduced into actin, demonstrating that filament severing is the essential function of cofilin in cells. This work reveals that site-specific interactions with cations serve a key regulatory function in actin filament fragmentation and dynamics.Actin polymerization powers the directed motility of eukaryotic cells and some pathogenic bacteria (1–3). Actin assembly also plays critical roles in endocytosis, cytokinesis, and establishment of cell polarity. Sustained motility requires filament disassembly and subunit recycling. The essential regulatory protein cofilin severs actin filaments (4–6), which accelerates actin network reorganization by increasing the concentration of filament ends available for subunit exchange (7).Cofilin binding alters the structure and mechanical properties of filaments, which effectively introduces local “defects” that compromise filament integrity and promote severing (5). Filaments with bound cofilin have altered twist (8, 9) and are more compliant in both bending and twisting than bare filaments (10–13). It has been suggested that deformations in filament shape promote fragmentation at or near regions of topological and mechanical discontinuities, such as boundaries between bare and cofilin-decorated segments along partially decorated filaments (5, 12, 14–18).Cations modulate actin filament structure and mechanical properties (19) and cofilin dissociates filament-associated cations (20), leading us to hypothesize that cation-binding interactions regulate filament severing by cofilin. Cations bind filaments at two discrete and specific sites positioned between adjacent subunits along the long-pitch helix of the filament (19, 21). These cation binding sites are referred to as “polymerization” and “stiffness” sites based on their roles in filament assembly and mechanics, respectively. These discrete sites bind both monovalent and divalent cations with a range of affinities (low millimolar for divalent and tens of millimolar for monovalent cations) (19, 21) but are predominantly occupied by Mg²⁺ and K⁺ under physiological conditions. Here we demonstrate that cation release from the stiffness site plays a central role in filament severing by vertebrate cofilin, both in vitro and in cells.     

Filopodial retraction force is generated by cortical actin dynamics and controlled by reversible tethering at the tip

Filopodia are dynamic, finger-like plasma membrane protrusions that sense the mechanical and chemical surroundings of the cell. Here, we show in epithelial cells that the dynamics of filopodial extension and retraction are determined by the difference between the actin polymerization rate at the tip and the retrograde flow at the base of the filopodium. Adhesion of a bead to the filopodial tip locally reduces actin polymerization and leads to retraction via retrograde flow, reminiscent of a process used by pathogens to invade cells. Using optical tweezers, we show that filopodial retraction occurs at a constant speed against counteracting forces up to 50 pN. Our measurements point toward retrograde flow in the cortex together with frictional coupling between the filopodial and cortical actin networks as the main retraction-force generator for filopodia. The force exerted by filopodial retraction, however, is limited by the connection between filopodial actin filaments and the membrane at the tip. Upon mechanical rupture of the tip connection, filopodia exert a passive retraction force of 15 pN via their plasma membrane. Transient reconnection at the tip allows filopodia to continuously probe their surroundings in a load-and-fail manner within a well-defined force range.Filopodia are actin-rich cell membrane protrusions, involved in processes as diverse as cell migration, wound closure, and cell invasion by pathogens (1–3). During cell migration, filopodia can exert forces on the substrate (4, 5) and act as precursors of focal adhesions (6–8). Filopodia initiate contacts during wound closure and contribute to dorsal closure of the fruit fly embryo in a zipper-like fashion (9–12). Viruses can hijack filopodia and filopodia-like cell–cell bridges to surf toward the cell body (13, 14). Filopodia from macrophages and epithelial cells actively pull pathogens bound to their tips (15–18). In all these examples filopodial retraction and retrograde force production are crucial. However, although filopodia formation and growth have been well studied (1–3), the mechanisms underlying their retraction are poorly understood.Filopodia show continuous rearward movement of their actin filaments in a process called “retrograde flow” (3, 19). In the lamellipodium, from which filopodia often emanate, the retrograde flow originates from actin treadmilling due to actin depolymerization at the rear and polymerization at the front of the lamellipodium. This retrograde flow is further amplified by the motor activity of myosins (20–23). In neurons, the filopodial shaft is deeply anchored in the growth cone and filopodial dynamics depends on the balance between actin polymerization at the filopodial tip and its retrograde flow (19). In other cell types actin depolymerization at the tip has been associated with retracting filopodia (24).Different contributions to filopodial force production during retraction can be considered. A connection between the filopodial tip and retracting actin filaments through transmembrane receptors such as integrins could transduce cortical forces applied on the actin shaft. In macrophages, force measurements on retracting filopodia suggested a major role for cortical myosins pulling on filopodial actin bundles (16). These measurements showed that retraction could be slowed down for forces below 20 pN. Applied forces higher than 20 pN inverted filopodial retraction of macrophages (25).Filopodial force production can also be due to membrane mechanics (26). Forces exerted by actin-free tubes extruded from the cell plasma membrane typically range between 5 pN and 30 pN (27). Membrane tension could drive filopodial retraction by exerting inward forces against the actin filaments. Moreover, filopodial actin filaments have been found disconnected from the membrane at the tip (28, 29), underlining the importance of membrane properties in filopodial mechanics. The contributions of membrane- and actin-based forces, as well as the mechanical links controlling force production during filopodial retraction, are still unclear.Here, we studied the retraction dynamics and the forces exerted by a single filopodium that is contacting an optically trapped bead at its tip. We found that filopodia retracted in association with a reduced actin polymerization at their tip at rates below those needed to compensate for the retrograde flow. The speed of filopodial retraction was only marginally affected by counteracting forces up to 50 pN, suggesting that the driving forces for retraction were not limiting within this range. We argue that actin treadmilling in the cell cortex, that functions far from its stall regime, transduces inward forces to the filopodial actin shaft at the base via high friction. In addition we found that filopodia can exert passive inward forces of 15 pN by using cell membrane-based forces. External counterforces that are only 5 pN higher than the membrane force can lead to rupture of connections between the actin shaft and the membrane at the filopodial tip. These weak contacts at the tip define the maximal pulling force of filopodia and allow cytoskeletal inward forces to operate only for short time intervals (<25 s). We found that the mechanical disconnection between membrane and actin filaments is only transient as actin dynamics at the tip are altered after disconnection. A continuous load-and-fail behavior allows thus tip-bound filopodia to probe the mechanics of their environment.     

Helical buckling of actin inside filopodia generates traction

Cells can interact with their surroundings via filopodia, which are membrane protrusions that extend beyond the cell body. Filopodia are essential during dynamic cellular processes like motility, invasion, and cell–cell communication. Filopodia contain cross-linked actin filaments, attached to the surrounding cell membrane via protein linkers such as integrins. These actin filaments are thought to play a pivotal role in force transduction, bending, and rotation. We investigated whether, and how, actin within filopodia is responsible for filopodia dynamics by conducting simultaneous force spectroscopy and confocal imaging of F-actin in membrane protrusions. The actin shaft was observed to periodically undergo helical coiling and rotational motion, which occurred simultaneously with retrograde movement of actin inside the filopodium. The cells were found to retract beads attached to the filopodial tip, and retraction was found to correlate with rotation and coiling of the actin shaft. These results suggest a previously unidentified mechanism by which a cell can use rotation of the filopodial actin shaft to induce coiling and hence axial shortening of the filopodial actin bundle.Tubular membrane remodeling driven by the actin cytoskeleton plays a major role in both pathogenesis and in a healthy immune response. For instance, invadopodia, podosomes, and filopodia are crucial for invasion and migration of cells (1). Filopodia are thin (100 to 300 nm), tube-like, actin-rich structures that function as “antennae” or “tentacles” that cells use to probe and interact with their microenvironment (2–4). Such structures have been studied in vitro using model systems where the point-like 3D contacts with the extracellular matrix (ECM) have been mimicked by using optically trapped dielectric particles, functionalized with relevant ligands. These model systems allow for mechanical and visual control over filopodial dynamics and have significantly advanced the understanding of cellular mechanosensing (5).Extraction of membrane tubes, using optical trapping, has been used to investigate mechanical properties of the membrane–cytoskeleton system (6–10), or membrane cholesterol content (11), and has revealed important insight into the mechanism that peripheral proteins use to shape membranes (12). Motivated by the pivotal role of F-actin in the mechanical behavior of filopodia and other cellular protrusions, special focus has been on revealing the presence of F-actin within extracted membrane tubes (1, 13–16). However, apart from a single study (9), fluorescent visualization of the F-actin was achieved by staining and fixation of the cells, and literature contains conflicting results regarding the presence or absence of actin in membrane tubes pulled from living cells (8, 11, 17).Filopodia in living cells have the ability to rotate and bend by a so-far unknown mechanism (13, 18–20). For instance, filopodia have been reported to exhibit sharp kinks in neuronal cells (21–23) and macrophages (6). These filopodial kinks have been observed both for surface-attached filopodia (23) as well as for filopodia that were free to rotate in three dimensions (6, 19).Other filaments such as DNA and bacterial flagella are common examples of structures that shorten and bend in response to torsional twist. Filopodia have previously been shown to have the ability to rotate by a mechanism that exists at their base (18, 19) and could have the ability to twist in presence of a frictional force. Rotation of the actin shaft results in friction with the surrounding filopodial membrane, and hence torsional energy can be transferred to the actin shaft. The myosin-Vb motor has been found to localize at the base of filopodia in neurite cells and to be critical for rotational movement of the filopodia; however, inhibition of myosin-Vc did not affect the rotation (19).Here, we reveal how actin filaments can simultaneously rotate and helically bend within cellular membrane tubes obtained by elongation of preexisting filopodia by an optically trapped bead. Simultaneous force measurements and confocal visualization reveal how the actin transduces a force as it rotates and retracts. After ∼100 s, the force exerted by the filopodium starts to exhibit pulling events reflecting transient contact between the actin and the tip region of the filopodium, which is attached to the optically trapped bead. We show that helical bending and rotation of the actin shaft (defined as the visible part of the actin) occurs simultaneously with movement of actin coils inside filopodia, which can occur concomitantly with a traction force exerted at the tip. The velocity of the coils depends on their location along the tube, thus indicating that retrograde flow is not the only mechanism driving the motion. Our data, and accompanying calculations, show that the rotation of the actin shaft, and the resulting torsional twist energy accumulated in the actin shaft, can contribute to shortening and bending of the actin shaft in conjunction with a retrograde flow.     

Random bursts determine dynamics of active filaments

Constituents of living or synthetic active matter have access to a local energy supply that serves to keep the system out of thermal equilibrium. The statistical properties of such fluctuating active systems differ from those of their equilibrium counterparts. Using the actin filament gliding assay as a model, we studied how nonthermal distributions emerge in active matter. We found that the basic mechanism involves the interplay between local and random injection of energy, acting as an analog of a thermal heat bath, and nonequilibrium energy dissipation processes associated with sudden jump-like changes in the system’s dynamic variables. We show here how such a mechanism leads to a nonthermal distribution of filament curvatures with a non-Gaussian shape. The experimental curvature statistics and filament relaxation dynamics are reproduced quantitatively by stochastic computer simulations and a simple kinetic model.In active systems, perpetual local energy input prevents relaxation into a thermal equilibrium state (1–3). Examples are living matter (4–10) or appropriately reconstituted or synthetic model systems (11–17). It is widely accepted that nonthermal fluctuations play a crucial role for the dynamics of active systems (8, 9, 18–24) and may even cause an apparent violation of the fluctuation-dissipation theorem (11). The physical origin of the violation can be attributed to local tensile stresses generated by myosin minifilaments, as shown by rheological measurements of 3D actin networks consisting of myosin II, actin filaments, and cross-linkers (11). Although this study focused on how the macroscopic properties of the active filament network are altered with respect to its equilibrium counterpart, we consider how local stresses generated by motors mesoscopically affect the dynamics and the conformational statistics of individual filaments. To this end, we use the actin gliding assay (25, 26), which has become a paradigm of active systems. In this assay, actin filaments are moved by individual nonprocessive myosin motors, which are bound to a substrate. We find that motile filaments in this assay display a nonthermal distribution of curvatures with an exponential shape, which is essentially different from its equilibrium counterpart. Based on our observations, we were able to elucidate the origin of the nonthermal fluctuations in the gliding assay and introduce a mechanism that explains how nonthermal distributions may emerge in active matter systems. The mechanism relies on the interplay between local and random input of energy, acting as an analog of a thermal heat bath, and nonequilibrium energy dissipation processes due to sudden jump-like changes in the system’s dynamic variables. We perform stochastic simulations of the filament’s dynamics and provide a rationale drawn from kinetic theory. Both approaches quantitatively reproduce the experimental curvature distribution and correctly predict the relaxation dynamics of the active filament.     

Self-assembly of alkynylplatinum(II) terpyridine amphiphiles into nanostructures via steric control and metal–metal interactions

A series of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing the hydrophilic oligo(para-phenylene ethynylene) with two 3,6,9-trioxadec-1-yloxy chains was designed and synthesized. The mononuclear alkynylplatinum(II) terpyridine complex was found to display a very strong tendency toward the formation of supramolecular structures. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would lead to the formation of nanotubes or helical ribbons. These desirable nanostructures were found to be governed by the steric bulk on the platinum(II) terpyridine moieties, which modulates the directional metal−metal interactions and controls the formation of nanotubes or helical ribbons. Detailed analysis of temperature-dependent UV-visible absorption spectra of the nanostructured tubular aggregates also provided insights into the assembly mechanism and showed the role of metal−metal interactions in the cooperative supramolecular polymerization of the amphiphilic platinum(II) complexes.Square-planar d⁸ platinum(II) polypyridine complexes have long been known to exhibit intriguing spectroscopic and luminescence properties (1–54) as well as interesting solid-state polymorphism associated with metal−metal and π−π stacking interactions (1–14, 25). Earlier work by our group showed the first example, to our knowledge, of an alkynylplatinum(II) terpyridine system [Pt(tpy)(C ≡ CR)]⁺ that incorporates σ-donating and solubilizing alkynyl ligands together with the formation of Pt···Pt interactions to exhibit notable color changes and luminescence enhancements on solvent composition change (25) and polyelectrolyte addition (26). This approach has provided access to the alkynylplatinum(II) terpyridine and other related cyclometalated platinum(II) complexes, with functionalities that can self-assemble into metallogels (27–31), liquid crystals (32, 33), and other different molecular architectures, such as hairpin conformation (34), helices (35–38), nanostructures (39–45), and molecular tweezers (46, 47), as well as having a wide range of applications in molecular recognition (48–52), biomolecular labeling (48–52), and materials science (53, 54). Recently, metal-containing amphiphiles have also emerged as a building block for supramolecular architectures (42–44, 55–59). Their self-assembly has always been found to yield different molecular architectures with unprecedented complexity through the multiple noncovalent interactions on the introduction of external stimuli (42–44, 55–59).Helical architecture is one of the most exciting self-assembled morphologies because of the uniqueness for the functional and topological properties (60–69). Helical ribbons composed of amphiphiles, such as diacetylenic lipids, glutamates, and peptide-based amphiphiles, are often precursors for the growth of tubular structures on an increase in the width or the merging of the edges of ribbons (64, 65). Recently, the optimization of nanotube formation vs. helical nanostructures has aroused considerable interests and can be achieved through a fine interplay of the influence on the amphiphilic property of molecules (66), choice of counteranions (67, 68), or pH values of the media (69), which would govern the self-assembly of molecules into desirable aggregates of helical ribbons or nanotube scaffolds. However, a precise control of supramolecular morphology between helical ribbons and nanotubes remains challenging, particularly for the polycyclic aromatics in the field of molecular assembly (64–69). Oligo(para-phenylene ethynylene)s (OPEs) with solely π−π stacking interactions are well-recognized to self-assemble into supramolecular system of various nanostructures but rarely result in the formation of tubular scaffolds (70–73). In view of the rich photophysical properties of square-planar d⁸ platinum(II) systems and their propensity toward formation of directional Pt···Pt interactions in distinctive morphologies (27–31, 39–45), it is anticipated that such directional and noncovalent metal−metal interactions might be capable of directing or dictating molecular ordering and alignment to give desirable nanostructures of helical ribbons or nanotubes in a precise and controllable manner.Herein, we report the design and synthesis of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing hydrophilic OPEs with two 3,6,9-trioxadec-1-yloxy chains. The mononuclear alkynylplatinum(II) terpyridine complex with amphiphilic property is found to show a strong tendency toward the formation of supramolecular structures on diffusion of diethyl ether in dichloromethane or dimethyl sulfoxide (DMSO) solution. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would result in nanotubes or helical ribbons in the self-assembly process. To the best of our knowledge, this finding represents the first example of the utilization of the steric bulk of the moieties, which modulates the formation of directional metal−metal interactions to precisely control the formation of nanotubes or helical ribbons in the self-assembly process. Application of the nucleation–elongation model into this assembly process by UV-visible (UV-vis) absorption spectroscopic studies has elucidated the nature of the molecular self-assembly, and more importantly, it has revealed the role of metal−metal interactions in the formation of these two types of nanostructures.     

Rasip1 mediates Rap1 regulation of Rho in endothelial barrier function through ArhGAP29

Rap1 is a small GTPase regulating cell–cell adhesion, cell–matrix adhesion, and actin rearrangements, all processes dynamically coordinated during cell spreading and endothelial barrier function. Here, we identify the adaptor protein ras-interacting protein 1 (Rasip1) as a Rap1-effector involved in cell spreading and endothelial barrier function. Using Förster resonance energy transfer, we show that Rasip1 interacts with active Rap1 in a cellular context. Rasip1 mediates Rap1-induced cell spreading through its interaction partner Rho GTPase-activating protein 29 (ArhGAP29), a GTPase activating protein for Rho proteins. Accordingly, the Rap1–Rasip1 complex induces cell spreading by inhibiting Rho signaling. The Rasip1–ArhGAP29 pathway also functions in Rap1-mediated regulation of endothelial junctions, which controls endothelial barrier function. In this process, Rasip1 cooperates with its close relative ras-association and dilute domain-containing protein (Radil) to inhibit Rho-mediated stress fiber formation and induces junctional tightening. These results reveal an effector pathway for Rap1 in the modulation of Rho signaling and actin dynamics, through which Rap1 modulates endothelial barrier function.The small GTPase Rap1 regulates both integrin-mediated and cadherin-mediated adhesions. Rap1 can increase cell adhesion by inducing the allosteric activation and clustering of integrins, thereby increasing cell–extracellular matrix (ECM) adhesion (1–3). Upon cell–ECM engagement, Rap1 induces cell spreading, due to increased cell protrusion and decreased cell contraction, indicating changes in actin dynamics (4, 5). In addition, Rap1 regulates both epithelial and endothelial cell–cell adhesion (6–11). Particularly the role of Rap1 in controlling endothelial cell junctions is important, as weakening of the endothelial barrier can result in pathologies such as chronic inflammation, atherosclerosis, and vascular leakage (12–14). Activation of Rap1 in endothelial cells results in stabilization of junctions and consequently increased barrier function through the recruitment of β-catenin, resulting in stabilization of vascular endothelial (VE)–cadherin at cell–cell junctions (15–18) and rearrangements of the actin cytoskeleton (6, 7, 19–21). These rearrangements of the actin cytoskeleton include the disruption of radial stress fibers and the induction of cortical actin bundles, and consequently a switch from discontinuous, motile junctions into linear, stable junctions (6–8, 20). Rap1 achieves this at least in part by regulating Rho-signaling (6, 7, 10, 19, 20). The molecular mechanism of how Rap1 regulates Rho, however, remains largely elusive, although the Rap1-effector Krev interaction trapped protein 1 (Krit-1)/cerebral cavernous malformations 1 protein (CCM1) has been proposed to be involved (15, 16, 22).In this study, we identified a Rap1-signaling cascade, comprising ras-interacting protein 1 (Rasip1), ras-association and dilute domain-containing protein (Radil), and Rho GTPase-activating protein 29 (ArhGAP29), affecting both cell spreading and endothelial barrier function by regulating the Rho-signaling cascade.     

Actin-dependent vacuolar occupancy of the cell determines auxin-induced growth repression

The cytoskeleton is an early attribute of cellular life, and its main components are composed of conserved proteins. The actin cytoskeleton has a direct impact on the control of cell size in animal cells, but its mechanistic contribution to cellular growth in plants remains largely elusive. Here, we reveal a role of actin in regulating cell size in plants. The actin cytoskeleton shows proximity to vacuoles, and the phytohormone auxin not only controls the organization of actin filaments but also impacts vacuolar morphogenesis in an actin-dependent manner. Pharmacological and genetic interference with the actin–myosin system abolishes the effect of auxin on vacuoles and thus disrupts its negative influence on cellular growth. SEM-based 3D nanometer-resolution imaging of the vacuoles revealed that auxin controls the constriction and luminal size of the vacuole. We show that this actin-dependent mechanism controls the relative vacuolar occupancy of the cell, thus suggesting an unanticipated mechanism for cytosol homeostasis during cellular growth.Actin filaments and its myosin motor proteins control a multitude of diverse cellular processes in animal cells, such as muscle contraction, cell motility, as well as vesicle and organelle movements (1). In animals, actin has a strong impact on the regulation of cellular shape and thus on cell size (2). Unlike animal cells, plant cells are sheathed by shape-giving cell walls, rendering them largely immobile. Despite this difference, the plant actin cytoskeleton has a conserved function in vesicle trafficking and organelle movement (3). Compared with animals, the role of actin in controlling cell size in plants is not clear and remains to be addressed. The phytohormone auxin is a crucial regulator of cell-size control in plants (4). Several studies suggest that the plant-specific growth regulator auxin affects the actin cytoskeleton (5–10). These studies concentrated on the effect of auxin on cortical actin and its contribution to processes close to the plasma membrane, such as endocytosis and exocytosis (5–11). Here we show that the actin cytoskeleton also is required for auxin processes beyond the plasma membrane, contributing to vacuolar morphogenesis and consequently to the regulation of cell size in plants.     

Two-tiered coupling between flowing actin and immobilized N-cadherin/catenin complexes in neuronal growth cones

Neuronal growth cones move forward by dynamically connecting actin-based motility to substrate adhesion, but the mechanisms at the individual molecular level remain unclear. We cultured primary neurons on N-cadherin–coated micropatterned substrates, and imaged adhesion and cytoskeletal proteins at the ventral surface of growth cones using single particle tracking combined to photoactivated localization microscopy (sptPALM). We demonstrate transient interactions in the second time scale between flowing actin filaments and immobilized N-cadherin/catenin complexes, translating into a local reduction of the actin retrograde flow. Normal actin flow on micropatterns was rescued by expression of a dominant negative N-cadherin construct competing for the coupling between actin and endogenous N-cadherin. Fluorescence recovery after photobleaching (FRAP) experiments confirmed the differential kinetics of actin and N-cadherin, and further revealed a 20% actin population confined at N-cadherin micropatterns, contributing to local actin accumulation. Computer simulations with relevant kinetic parameters modeled N-cadherin and actin turnover well, validating this mechanism. Such a combination of short- and long-lived interactions between the motile actin network and spatially restricted adhesive complexes represents a two-tiered clutch mechanism likely to sustain dynamic environment sensing and provide the force necessary for growth cone migration.Growth cones are motile structures at the extremity of axons responsible for path finding and neurite extension during nervous system development and repair. Growth cones translate extracellular signals into directional migration through a coordinated regulation of cytoskeleton, adhesion, and membrane processes (1). At the cytoskeletal level, motility is generated by polarized actin treadmilling, which, together with myosin contraction, generates a continuous retrograde actin flow from the periphery to the base of growth cones (2–7). At the membrane level, adhesion proteins form dynamic bonds with immobilized extracellular ligands, allowing step-by-step locomotion (8).The molecular clutch model postulates that the mechanical coupling between ligand-bound transmembrane adhesion receptors and the actin flow allows traction forces to be transmitted to the substrate, resulting in local diminution of the retrograde flow and forward progression (9–11). Optical tweezers and flexible substrate experiments using microspheres coated with adhesion molecules revealed clutch-like mechanisms for integrins (12, 13), Ig cell adhesion molecules (14, 15), and cadherins (16, 17). However, the mechanism of clutch engagement at the individual molecular level remains elusive. For integrin-based adhesion, single-molecule tracking experiments suggested that talin and vinculin could switch between a state bound to flowing actin and a state bound to immobilized integrins (18, 19). For cadherin-based adhesion, biochemical experiments suggested that α-catenin could transit between being bound to actin or to the cadherin/β-catenin complex (20, 21), but a direct visualization of such behavior is lacking. In addition, vinculin, which can bind both actin and α-catenin, is recruited at cadherin-based intercellular junctions under mechanical force (22–24), but its dynamic behavior in growth cone migration is unknown.By combining spatially controlled adhesion in growth cones with single-molecule tracking, fluorescence recovery after photobleaching (FRAP) experiments, and computer simulations, we report that N-cadherin/α-catenin complexes, but not vinculin, are selectively trapped at N-cadherin micropatterns. In addition, 80% of actin molecules flow rearward, making transient pauses on the order of seconds with immobilized N-cadherin/α-catenin complexes, whereas 20% of confined actin molecules contribute to local actin enrichment. This association of short- and long-lasting individual bonds underlies the differential coupling between the actin motile machinery and substrate adhesions supporting growth cone migration.