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.     

Inverted formin 2 in focal adhesions promotes dorsal stress fiber and fibrillar adhesion formation to drive extracellular matrix assembly

Actin filaments and integrin-based focal adhesions (FAs) form integrated systems that mediate dynamic cell interactions with their environment or other cells during migration, the immune response, and tissue morphogenesis. How adhesion-associated actin structures obtain their functional specificity is unclear. Here we show that the formin-family actin nucleator, inverted formin 2 (INF2), localizes specifically to FAs and dorsal stress fibers (SFs) in fibroblasts. High-resolution fluorescence microscopy and manipulation of INF2 levels in cells indicate that INF2 plays a critical role at the SF–FA junction by promoting actin polymerization via free barbed end generation and centripetal elongation of an FA-associated actin bundle to form dorsal SF. INF2 assembles into FAs during maturation rather than during their initial generation, and once there, acts to promote rapid FA elongation and maturation into tensin-containing fibrillar FAs in the cell center. We show that INF2 is required for fibroblasts to organize fibronectin into matrix fibers and ultimately 3D matrices. Collectively our results indicate an important role for the formin INF2 in specifying the function of fibrillar FAs through its ability to generate dorsal SFs. Thus, dorsal SFs and fibrillar FAs form a specific class of integrated adhesion-associated actin structure in fibroblasts that mediates generation and remodeling of ECM.The dynamic connection between the forces generated in the actomyosin cytoskeleton and integrin-mediated focal adhesions (FAs) to the extracellular matrix (ECM) is essential for many physiological processes including cell migration, vascular formation and function, the immune response, and tissue morphogenesis. These diverse functions are mediated by distinct cellular structures including protruding lamellipodia containing nascent FAs that mediate haptotaxis (1), ventral adhesive actin waves that mediate leukocyte transmigration through endothelia (2, 3), and stress fibers (SFs) and FAs that drive fibrillarization of ECM in developing embryos (4, 5). The coordination and interdependence of actin and integrin-based adhesion in these specialized cellular structures are rooted in their biochemical interdependence. Activation of integrins to their high-affinity ECM binding state requires the actin cytoskeleton (6). In turn, integrin engagement with ECM induces signaling that mediates actin polymerization and contractility downstream of Rho GTPases (6, 7). ECM-engaged integrins also affect cytoskeletal organization by physically linking the contractile actomyosin system to extracellular anchorage points (7). Thus, adhesion-associated actin structures are integrated systems that mediate cellular functions requiring coordination of intracellular cytoskeletal forces with ECM binding.Mesenchymal cells generally possess two main types of adhesion-associated actin structures: protruding lamellipodia containing nascent FAs at the cell edge and linear actin bundles in the cell body connected to FAs. Compared with architecturally invariant lamellipodia, adhesion-associated actin bundle structures, including filopodia, the perinuclear actin cap/transmembrane actin-associated nuclear lines, trailing edge bundles, and dorsal SFs, are more diverse in their morphology and less well understood in their architecture and function (8–10). The most-studied actin bundle structure is perhaps dorsal SFs, noncontractile bundles associated at one end with a ventral FA near the cell edge and that extend radially toward the cell center and join with dorsal actin arcs on their other end. How the functional specificity of dorsal SFs is generated apart from the many other distinct adhesion-associated actin bundle structures is not well understood.The functional specificity of adhesion-associated actin structures could be generated either on the adhesion side by compositional differences in FA proteins or on the actin side by differences in the nucleation mechanism and actin binding proteins. On the adhesion side, it is well known that different integrin family members bind distinct types of ECM (11, 12). However, cells adhered to different ECMs all form common structures including lamellipodia, filopodia, and multiple types of SFs. In addition to different integrins, FA function could be regulated by the process of “maturation” in which FAs undergo stereotypical dynamic changes in composition and morphology driven by actomyosin-mediated cellular tension (13, 14). Nascent FAs contain integrins, focal adhesion kinase (FAK), a-actinin, and paxillin (13, 15). When tension is applied, nascent FAs grow and recruit hundreds of proteins, including talin, vinculin, and zyxin (16). These mature FAs then either disassemble or further mature into tensin-containing fibrillar FAs that are responsible for fibronectin fibrillogenesis (17). Thus, the changes in FA size and protein content that accompany FA maturation could give rise to functional specialization of adhesion/actin systems.On the other hand, actin filaments in migrating cells are generated by two main classes of nucleators: the Arp2/3 complex and formins (18). Different nucleating proteins generate different actin organization and geometries, which could in turn dictate functional specificity of adhesions. Arp2/3 forms the branched network in lamellipodia and is thought to be linked to nascent FAs through interaction with FAK (19–21) or vinculin (22). The formin family of actin nucleators, which generates linear actin bundles (23), is more diverse, although formins share a common actin assembly core domain (24), (25). Recent work has begun to ascribe the generation of particular actin structures to some of the 15 formins in mammalian cells, particularly members of the diaphanous family and FHOD1 (26–29). Specifically regarding dorsal SFs, evidence points strongly to polymerization by a formin family member (23, 30–32) but no formin has ever been localized to these SFs or their associated FAs in motile cells. Thus, although formins are clearly critical for forming distinct actin structures, whether they cooperate with FA proteins to specify the function of adhesion-associated actin structures in the cell is unclear.We hypothesized that inverted formin 2 (INF2), found in our recent FA proteome (33), may play a critical role in the formation and functional specificity of adhesion-associated actin structures. INF2 is expressed in cells in two isoforms, one containing a membrane-targeting CAAX-motif that plays a role in mitochondrial fission (34) and a non-CAAX isoform whose function is not well characterized. INF2 is an unusual formin insofar as it contains, in addition to the FH1–FH2 domains that polymerize actin, a WH2-like domain at the C terminus (35) that binds actin monomers to regulate autoinhibition, and also mediates filament severing (35, 36). INF2 also interacts with and inhibits members of the diaphanous family of formin proteins (37). INF2 therefore could have multiple possible roles at FAs in local modulation of actin.Here we explore the role of INF2 in mouse embryonic fibroblasts (MEFs). We find for the first time to our knowledge strong localization of an endogenous formin to FAs at the distal tips of dorsal SFs where it is required for actin polymerization at FAs to form dorsal SFs. We show that INF2 plays a role in controlling morphological, but not compositional maturation of FAs. Strikingly, INF2 is responsible for the formation of one specific class of FAs, the fibrillar FAs that organize the ECM; disruption of INF2 leads to defects in ECM fibrillogenesis. Thus, our study demonstrates that INF2 mediates the formation of dorsal SFs and fibrillar FAs, which together comprise a specific integrated adhesion-associated actin structure responsible for the fibrillogenesis of ECM by fibroblasts.     

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.     

Capping protein regulatory cycle driven by CARMIL and V-1 may promote actin network assembly at protruding edges

Although capping protein (CP) terminates actin filament elongation, it promotes Arp2/3-dependent actin network assembly and accelerates actin-based motility both in vitro and in vivo. In vitro, capping protein Arp2/3 myosin I linker (CARMIL) antagonizes CP by reducing its affinity for the barbed end and by uncapping CP-capped filaments, whereas the protein V-1/myotrophin sequesters CP in an inactive complex. Previous work showed that CARMIL can readily retrieve CP from the CP:V-1 complex, thereby converting inactive CP into a version with moderate affinity for the barbed end. Here we further clarify the mechanism of this exchange reaction, and we demonstrate that the CP:CARMIL complex created by complex exchange slows the rate of barbed-end elongation by rapidly associating with, and dissociating from, the barbed end. Importantly, the cellular concentrations of V-1 and CP determined here argue that most CP is sequestered by V-1 at steady state in vivo. Finally, we show that CARMIL is recruited to the plasma membrane and only at cell edges undergoing active protrusion. Assuming that CARMIL is active only at this location, our data argue that a large pool of freely diffusing, inactive CP (CP:V-1) feeds, via CARMIL-driven complex exchange, the formation of weak-capping complexes (CP:CARMIL) at the plasma membrane of protruding edges. In vivo, therefore, CARMIL should promote Arp2/3-dependent actin network assembly at the leading edge by promoting barbed-end capping there.Actin assembly at the interface between the plasma membrane and the cytoplasm is driven by the recruitment of proteins that promote the nucleation of new actin filaments (1). A key nucleating factor driving actin assembly at the leading edge of crawling cells is the Arp2/3 complex, which creates the branched actin networks that comprise lamellipodia (2, 3). Capping protein (CP), which binds the barbed end of the actin filament with very high affinity (∼0.1 nM) to halt elongation (4), is an essential component of Arp2/3-generated branched actin networks (5). Paradoxically, although CP terminates filament elongation, it promotes Arp2/3-dependent actin network assembly and actin-based motility both in vitro and in vivo by increasing the frequency of Arp2/3-dependent nucleation (6–10). Factors that regulate the activity of CP therefore should play important roles in actin assembly and cell motility. One such factor may be the large scaffold protein CARMIL (capping protein Arp2/3 myosin I linker) (11), which contains a region of ∼75 residues that binds CP very tightly (K_d ∼1 nM) (12, 13). This region, which we named CAH3 for CARMIL Homology Domain 3, antagonizes the function of CP in vitro in two ways. First, by binding to free CP, CAH3 reduces CP’s affinity for the barbed end from ∼0.1 to ∼50 nM, thereby creating a weak barbed-end capping complex (CP:CAH3) (12, 13). Second, by binding to CP already present on the barbed end, CAH3 accelerates by ∼300-fold the rate of dissociation of CP from the end; i.e., CAH3 rapidly uncaps CP-capped filaments (12–14).Another potentially important regulator of CP is the protein V-1, also known as myotrophin (15, 16). This 13 kDa, ankyrin-repeat protein binds CP with an affinity of ∼20 nM, creating a 1:1 complex (CP:V-1) that has no affinity for the barbed end (16–18). V-1 has the potential, therefore, to influence polymerization by reducing the extent of barbed-end capping.Full understanding of CP regulation will require an in-depth understanding of how the activities of CARMIL and V-1 are coordinated. Relevant to this, Takeda et al. (17) showed that CAH3-like peptides readily convert the CP:V-1 complex into the CP:CAH3 complex. In terms of the mechanism of this exchange reaction, they postulated that the binding of CAH3 to the CP:V-1 complex results in the transient formation of a CP:V-1:CAH3 ternary complex. A subsequent allosteric change in CP caused by ternary complex formation then drives V-1 dissociation, yielding the CP:CAH3 complex. Importantly, the binding sites on the surface of CP for CAH3 and V-1 are nonoverlapping, consistent with the possibility that these three proteins form a ternary complex (17–21).The first goal of this study was to define precisely how CAH3 catalyzes the exchange of CP from its sequestered state (CP:V-1) into its weak barbed-end–binding state (CP:CAH3). This effort focused on providing direct evidence for the existence of the ternary complex and on determining the relative contributions of allostery versus competitive binding to complex exchange. The second goal was to determine the effect of complex exchange on the rate of barbed-end elongation in vitro, which must be understood to predict the effect of complex exchange on actin assembly in vivo. The third goal was to measure the cellular concentrations of V-1 and CP, as these values are required to gauge the likely significance of the complex exchange reaction in vivo. Finally, the fourth goal was to further define the spatial features and temporal dynamics of CARMIL’s leading-edge localization (11, 12, 22, 23), as this should further pinpoint both the site and the timing of complex exchange. Specifically, we sought to characterize CARMIL’s leading-edge localization relative to other key leading-edge molecules and to the plasma membrane and to determine if CARMIL’s leading-edge accumulation correlates with edge dynamics, i.e., protrusion versus retraction. Overall, our results support a model in which the recruitment of CARMIL to the plasma membrane specifically at sites of active polymerization and edge protrusion drives the exchange of abundant, freely diffusing, inactive CP:V-1 complexes into membrane-bound CP:CARMIL complexes that cap the barbed end weakly. This CARMIL-driven exchange reaction should therefore serve to promote Arp2/3-dependent actin network formation at protruding edges by promoting barbed-end capping there.     

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.     

Actin-related protein2/3 complex regulates tight junctions and terminal differentiation to promote epidermal barrier formation

The epidermis provides an essential seal from the external environment and retains fluids within the body. To form an effective barrier, cells in the epidermis must form tight junctions and terminally differentiate into cornified envelopes. Here, we demonstrate that the branched actin nucleator, the actin-related protein (Arp)2/3 complex, is unexpectedly required for both these activities. Loss of the ArpC3 subunit of the Arp2/3 complex resulted in minimal changes in the morphogenesis and architecture of this stratified squamous epithelium, but resulted in profound defects in its physiology. Mutant embryos did not develop an effective barrier to the external environment and died within hours of birth. We discovered two underlying causes for these effects. First, ArpC3 was essential for robust assembly and function of tight junctions, specialized cell–cell adhesions that restrict water loss in the epidermis. Second, there were defects in differentiation of the epidermis and the production of cornified envelopes, structures essential for barrier activity. Underlying this defect, we found that YAP was inappropriately active not only in the ArpC3 mutant tissue, but also in cultured cells. Inhibition of YAP activity rescued the differentiation and barrier defects caused by loss of ArpC3. These results demonstrate previously unappreciated roles for the Arp2/3 complex and highlight the functions of branched actin networks in a complex tissue.The epidermis is a stratified squamous epithelium that forms a barrier between us and our environment. Although the 3D architecture of this tissue is required for its function, we have only a rudimentary understanding of how different cytoskeletal structures function to control tissue organization and physiology. The actin cytoskeleton is a dynamic structural component of the cell that is necessary for cell shape, migration, and adhesion. Because of its many roles, actin is essential and therefore cannot be studied by direct loss-of-function approaches. However, many proteins regulate the assembly, bundling, cross-linking, capping, severing, and disassembly of F-actin to generate diverse cytoskeletal structures. These include proteins that promote the nucleation of new actin filaments, such as the actin-related protein (Arp)2/3 complex, formins, and proteins with multiple G-actin binding motifs like cordon-bleu and Spire (1). Of these, the Arp2/3 complex is unique in that it promotes the formation of branched actin networks.The mechanism and regulation of Arp2/3 complex-induced actin assembly has been extensively studied biochemically (2, 3). Cryo-EM and X-ray crystallography have yielded structural insights into the organization and interaction of the complex with actin filaments (4, 5). In addition, work in cultured cells has uncovered many Arp2/3 complex-dependent processes, including lamellipodia formation, efficient cell migration, endocytosis, vesicle trafficking, and adherens junction formation (6–10). Although the biochemistry and cell biology of the Arp2/3 complex have been well studied, most of our understanding of its role in intact tissues comes from invertebrate model systems. For example, the Arp2/3 complex is required for cell polarity and gastrulation in Caenorhabditis elegans embryos (11, 12). In Drosophila, the Arp2/3 complex plays roles in myoblast fusion, ring canal expansion, adherens junction formation, Notch signaling, and bristle development (6, 13, 14).The functions of the Arp2/3 complex in intact tissues during mammalian development and homeostasis have only begun to be addressed. Loss of the Arp2/3 complex globally results in early lethality at the blastocyst stage in mice (15). In the mouse oocyte, the Arp2/3 complex is required for asymmetric cell division through positioning of the mitotic spindle (16). Finally, specific deletion of ArpC3 from postmitotic neurons resulted in delayed defects in dendritic spine morphology and schizophrenia-like phenotypes (17). These studies highlight that loss of the Arp2/3 complex in a tissue-specific manner can uncover unexpected roles for branched actin networks.We wished to determine the roles of the Arp2/3 complex in the development and function of the epidermis, a stratified squamous epithelium. This tissue contains polarized cuboidal progenitor cells that undergo asymmetric cell divisions to form a multilayered, differentiated tissue that acts as a barrier to the external environment. Full barrier activity requires tight junction formation in the granular layer of the epidermis and the differentiation of cells into cornified envelopes that form the outer layer of the skin. Epidermal cells form robust adhesions to each other and to the underlying basement membrane and provide a physiological tissue context to analyze Arp2/3 function. We therefore generated mice with epidermal ablation of ArpC3, a peripheral subunit of the Arp2/3 complex required for robust F-actin nucleation. We find that ArpC3 is not required for cell shape, cell adhesion, or proper development of the 3D architecture of the epidermis. Unexpectedly, however, we find roles for ArpC3 in tight junction assembly and function and in YAP-mediated differentiation of the epidermis, highlighting specific roles for core cell biological machinery in tissue physiology.     

Lytic immune synapse function requires filamentous actin deconstruction by Coronin 1A

Lytic immune effector function depends upon directed secretion of cytolytic granules at the immunological synapse (IS) and requires dynamic rearrangement of filamentous (F)-actin. Coronin 1A (Coro1A) is the hematopoietic-specific member of the Coronin family of actin regulators that promote F-actin disassembly. Here, we show that Coro1A is required for natural killer (NK) cell cytotoxic function in two human NK cell lines and ex vivo cells from a Coro1A-deficient patient. Using superresolution nanoscopy to probe the IS, we demonstrate that Coro1A promotes the deconstruction of F-actin density that facilitates effective delivery of lytic granules to the IS. Thus, we show, for the first time to our knowledge, a critical role for F-actin deconstruction in cytotoxic function and immunological secretion and identify Coro1A as its mediator.Natural killer (NK) cell cytotoxicity is a finely controlled process that integrates signals from activating and inhibitory receptors to eliminate virally infected and tumorigenic cells sensitively and specifically. The importance for NK cells in immune function is underscored by the severe virus infections and malignancies suffered by patients with NK cell deficiency (1). A dynamic filamentous (F)-actin cytoskeleton is required for NK cell cytotoxicity because disruption of F-actin polymerization by pharmacological inhibitors or mutation of actin-nucleating factors results in impaired NK cell function (2–5). Actin nucleators, such as actin-related proteins 2 and 3 complex (Arp2/3), Wiskott–Aldrich syndrome protein (WASp), WIP, DOCK8, and WAVE2, serve well-defined critical roles in the formation and function of the NK cell immunological synapse (IS) (4–10).Killing of a susceptible target follows tightly regulated steps of NK cell immune synapse formation and lytic granule exocytosis (3). Although cortical F-actin has long been considered a barrier to exocytosis of granule-like organelles (11) in some cell types, ligation of NK cell activating and adhesion receptors results in the formation of conduits in F-actin that permit and actually facilitate NK cell degranulation (12–14). This finding suggests that fine regulation and deconstruction of the synaptic F-actin meshwork is required for the formation of granule-permissive–sized clearances (12).Coronin 1A (Coro1A) is the hematopoietic cell-specific isoform of the highly conserved Coronin family of actin regulators. Coronins contain a series of WD-repeat domains that form an F-actin–binding β-propeller domain, and thus bind F-actin directly (15–17). In addition, Coro1A binds to and inhibits the Arp2/3 complex (17, 18) required for actin branching and can enhance the activity of cofilin to promote actin disassembly in in vitro reconstituted systems (19–21). Coro1A localizes with actin-rich structures in immune cells, including phagocytic cups in neutrophils and macrophages, and at the leading edge of T cells (22–25). T cells from Coronin 1^−/− mice have defects in migration and cell survival attributed to impaired T-cell receptor signaling, Ca²⁺ flux, Rac activation, and subcellular Arp2/3 localization (26–28). Mutations in Coro1A lead to T⁻B⁺NK⁺ combined immunodeficiency and susceptibility to severe viral infections, including life-threatening varicella infection and EBV-driven lymphoproliferation (26, 29, 30).By manipulating expression of Coro1A in human NK cells, we show that Coro1A is required for cytotoxic function. Using superresolution nanoscopy, we define a requirement for Coro1A in F-actin deconstruction and subsequent delivery of lytic granules to the synaptic membrane. In addition, we have specifically evaluated cytotoxic function in a Coro1A-deficient patient and find that NK cell function is severely impaired. Further, we demonstrate the same F-actin structural defect in patient cells as in two Coro1A-deficient cell lines. Thus, with superresolution imaging, we identify, for the first time to our knowledge, a critical role for actin deconstruction in immunity and human host defense.     

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.     

From the Cover: Cortical instability drives periodic supracellular actin pattern formation in epithelial tubes

An essential question of morphogenesis is how patterns arise without preexisting positional information, as inspired by Turing. In the past few years, cytoskeletal flows in the cell cortex have been identified as a key mechanism of molecular patterning at the subcellular level. Theoretical and in vitro studies have suggested that biological polymers such as actomyosin gels have the property to self-organize, but the applicability of this concept in an in vivo setting remains unclear. Here, we report that the regular spacing pattern of supracellular actin rings in the Drosophila tracheal tubule is governed by a self-organizing principle. We propose a simple biophysical model where pattern formation arises from the interplay of myosin contractility and actin turnover. We validate the hypotheses of the model using photobleaching experiments and report that the formation of actin rings is contractility dependent. Moreover, genetic and pharmacological perturbations of the physical properties of the actomyosin gel modify the spacing of the pattern, as the model predicted. In addition, our model posited a role of cortical friction in stabilizing the spacing pattern of actin rings. Consistently, genetic depletion of apical extracellular matrix caused strikingly dynamic movements of actin rings, mirroring our model prediction of a transition from steady to chaotic actin patterns at low cortical friction. Our results therefore demonstrate quantitatively that a hydrodynamical instability of the actin cortex can trigger regular pattern formation and drive morphogenesis in an in vivo setting.Self-organization is one of the principal mechanisms of biological pattern formation at the molecular, cellular, and tissue scale. Although the pioneering work of Turing (1) has suggested reaction–diffusion as a generic route toward pattern generation (2), a concrete biomolecular or mechanical understanding of how this might occur in vivo remains elusive, except in a few specific cases (3–5). For instance, Kondo and coworkers (6) demonstrated that pigment patterning on the skin of the Pomocanthus imperator can be understood quantitatively from the simple attraction–repulsion kinetics of two cell types.At the cellular level, active structures, such as the cytoskeleton, are generically expected to display a large variety of structures from a theoretical perspective (7–12), many of which have been reproduced in elegant in vitro studies (13–15). In the case of actomyosin gels, the contractile stresses arising from molecular motors have been shown to create large actin flows that can reorganize the cortex (16, 17). Because actin filaments and motors are “self-advected,” or transported, by their own flow (18), there is a self-reinforcing loop in gel density, capable of creating patterns. Nevertheless, most theoretical studies do not consider the cross-effects of polymerization and diffusion, which resist pattern formation. Interestingly, in the past years, several groups have reported in vivo examples of actin patterns: mammalian axons (19), Caenorhabditis elegans embryo (20), and Drosophila trachea (21) are all cellular cylinders that display a regular array of concentric actin rings on their cortex.In this article, we study the example of ring formation in the Drosophila trachea and propose a generic mechanism for stable actin pattern formation, arising from the interplay of actin turnover and myosin activity. The model makes clear predictions, which we test through fly genetics and drug experiments.     

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.     

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.     

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.