Cell-sized liposomes reveal how actomyosin cortical tension drives shape change

Animal cells actively generate contractile stress in the actin cortex, a thin actin network beneath the cell membrane, to facilitate shape changes during processes like cytokinesis and motility. On the microscopic scale, this stress is generated by myosin molecular motors, which bind to actin cytoskeletal filaments and use chemical energy to exert pulling forces. To decipher the physical basis for the regulation of cell shape changes, here, we use a cell-like system with a cortex anchored to the outside or inside of a liposome membrane. This system enables us to dissect the interplay between motor pulling forces, cortex–membrane anchoring, and network connectivity. We show that cortices on the outside of liposomes either spontaneously rupture and relax built-up mechanical stress by peeling away around the liposome or actively compress and crush the liposome. The decision between peeling and crushing depends on the cortical tension determined by the amount of motors and also on the connectivity of the cortex and its attachment to the membrane. Membrane anchoring strongly affects the morphology of cortex contraction inside liposomes: cortices contract inward when weakly attached, whereas they contract toward the membrane when strongly attached. We propose a physical model based on a balance of active tension and mechanical resistance to rupture. Our findings show how membrane attachment and network connectivity are able to regulate actin cortex remodeling and membrane-shape changes for cell polarization.Animal cells constantly adapt their shape as they move and divide. Events like cytokinesis (1) and motility (2) require concerted remodeling of the actin cytoskeleton and the plasma membrane. A crucial cell module that drives cellular shape changes is the actomyosin cortex beneath the cell membrane, which produces contractile forces that squeeze the cell forward during migration or constrict it during division. The cell cortex is a thin actin network of thickness ∼0.2 µm (3), which is tightly attached to the plasma membrane (4, 5). On the molecular scale, contractile forces are generated by myosin II motor proteins, associated into bipolar filaments (6), which use adenosine 5′-triphosphate (ATP) to exert pulling forces on actin filaments.To drive cell-shape changes, these forces must be communicated to the plasma membrane, which requires membrane–actin attachment (7–9) through, for example, proteins from the ERM (ezrin, radixin, moesin) family (10–13). Several in vivo studies provide evidence that cortex–membrane attachment strongly influences contractile processes. For instance, blebbing at the cell poles due to transient detachment of the cortex from the membrane serves to release excess cortical tension during cytokinesis, which is crucial to achieve proper division (14). In early Caenorhabditis elegans embryos, myosin-driven contraction drives long-range cortical flow of actin along the membrane, which is crucial to establish cell polarity and asymmetric cell division (15). It is increasingly recognized that cortex remodeling and cell-shape changes depend on a balance between the active forces generated by motors and the passive forces originating from cortex (visco)elasticity and cortex–membrane adhesion (16). However, the microscopic basis of this force balance remains unclear. It is difficult to resolve this question directly in cells, because manipulation by drugs or genetic methods can also affect the dynamics of the underlying cytoskeleton via specific signaling pathways (17).An alternative approach is to use either cell-free extracts (18, 19) or systems reconstituted from a minimal set of purified cellular components (20). These approaches have been successfully used to show that contractility of bulk actomyosin networks depends on the kinetic parameters of the motors (21–23), as well as on the presence of actin cross-linkers that allow build-up of stress (24, 25). A recent study of 2D actomyosin networks attached to flat model biomembranes showed that actin–membrane anchoring also influences cortex contractility (26). However, although this biomimetic assay can address actin–membrane adhesion, it cannot address the influence of membrane deformability on cortex remodeling and shape change.Here, we reproduce active contractility in a cell-like system, where actomyosin cortical networks are anchored to a liposome membrane. Cortices linked to the outer leaflet of the liposome membrane mimic the cell cortex as well as intracellular organelles like endosomes (27). Low linkage or cross-link density promote active cortex rupture, which breaks the symmetry and causes passive elastic retraction of the network to one pole of the liposome. In contrast, high membrane anchorage and high network cross-link density promote active compression of the liposomes by cortex contraction. We further show that when the cortices are instead connected to the inner leaflet of the liposome membrane, actin–membrane linkage strongly biases the directionality of cortex contraction. Our results shed light on the physical mechanisms that control contraction events during cell division, motility and endosome-shape changes.     

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.     

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.     

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.     

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.     

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.     

Structural interactions of a voltage sensor toxin with lipid membranes

Protein toxins from tarantula venom alter the activity of diverse ion channel proteins, including voltage, stretch, and ligand-activated cation channels. Although tarantula toxins have been shown to partition into membranes, and the membrane is thought to play an important role in their activity, the structural interactions between these toxins and lipid membranes are poorly understood. Here, we use solid-state NMR and neutron diffraction to investigate the interactions between a voltage sensor toxin (VSTx1) and lipid membranes, with the goal of localizing the toxin in the membrane and determining its influence on membrane structure. Our results demonstrate that VSTx1 localizes to the headgroup region of lipid membranes and produces a thinning of the bilayer. The toxin orients such that many basic residues are in the aqueous phase, all three Trp residues adopt interfacial positions, and several hydrophobic residues are within the membrane interior. One remarkable feature of this preferred orientation is that the surface of the toxin that mediates binding to voltage sensors is ideally positioned within the lipid bilayer to favor complex formation between the toxin and the voltage sensor.Protein toxins from venomous organisms have been invaluable tools for studying the ion channel proteins they target. For example, in the case of voltage-activated potassium (Kv) channels, pore-blocking scorpion toxins were used to identify the pore-forming region of the channel (1, 2), and gating modifier tarantula toxins that bind to S1–S4 voltage-sensing domains have helped to identify structural motifs that move at the protein–lipid interface (3–5). In many instances, these toxin–channel interactions are highly specific, allowing them to be used in target validation and drug development (6–8).Tarantula toxins are a particularly interesting class of protein toxins that have been found to target all three families of voltage-activated cation channels (3, 9–12), stretch-activated cation channels (13–15), as well as ligand-gated ion channels as diverse as acid-sensing ion channels (ASIC) (16–21) and transient receptor potential (TRP) channels (22, 23). The tarantula toxins targeting these ion channels belong to the inhibitor cystine knot (ICK) family of venom toxins that are stabilized by three disulfide bonds at the core of the molecule (16, 17, 24–31). Although conventional tarantula toxins vary in length from 30 to 40 aa and contain one ICK motif, the recently discovered double-knot toxin (DkTx) that specifically targets TRPV1 channels contains two separable lobes, each containing its own ICK motif (22, 23).One unifying feature of all tarantula toxins studied thus far is that they act on ion channels by modifying the gating properties of the channel. The best studied of these are the tarantula toxins targeting voltage-activated cation channels, where the toxins bind to the S3b–S4 voltage sensor paddle motif (5, 32–36), a helix-turn-helix motif within S1–S4 voltage-sensing domains that moves in response to changes in membrane voltage (37–41). Toxins binding to S3b–S4 motifs can influence voltage sensor activation, opening and closing of the pore, or the process of inactivation (4, 5, 36, 42–46). The tarantula toxin PcTx1 can promote opening of ASIC channels at neutral pH (16, 18), and DkTx opens TRPV1 in the absence of other stimuli (22, 23), suggesting that these toxin stabilize open states of their target channels.For many of these tarantula toxins, the lipid membrane plays a key role in the mechanism of inhibition. Strong membrane partitioning has been demonstrated for a range of toxins targeting S1–S4 domains in voltage-activated channels (27, 44, 47–50), and for GsMTx4 (14, 50), a tarantula toxin that inhibits opening of stretch-activated cation channels in astrocytes, as well as the cloned stretch-activated Piezo1 channel (13, 15). In experiments on stretch-activated channels, both the d- and l-enantiomers of GsMTx4 are active (14, 50), implying that the toxin may not bind directly to the channel. In addition, both forms of the toxin alter the conductance and lifetimes of gramicidin channels (14), suggesting that the toxin inhibits stretch-activated channels by perturbing the interface between the membrane and the channel. In the case of Kv channels, the S1–S4 domains are embedded in the lipid bilayer and interact intimately with lipids (48, 51, 52) and modification in the lipid composition can dramatically alter gating of the channel (48, 53–56). In one study on the gating of the Kv2.1/Kv1.2 paddle chimera (53), the tarantula toxin VSTx1 was proposed to inhibit Kv channels by modifying the forces acting between the channel and the membrane. Although these studies implicate a key role for the membrane in the activity of Kv and stretch-activated channels, and for the action of tarantula toxins, the influence of the toxin on membrane structure and dynamics have not been directly examined. The goal of the present study was to localize a tarantula toxin in membranes using structural approaches and to investigate the influence of the toxin on the structure of the lipid bilayer.     

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.     

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.     

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.     

A RhoA and Rnd3 cycle regulates actin reassembly during membrane blebbing

The actin cytoskeleton usually lies beneath the plasma membrane. When the membrane-associated actin cytoskeleton is transiently disrupted or the intracellular pressure is increased, the plasma membrane detaches from the cortex and protrudes. Such protruded membrane regions are called blebs. However, the molecular mechanisms underlying membrane blebbing are poorly understood. This study revealed that epidermal growth factor receptor kinase substrate 8 (Eps8) and ezrin are important regulators of rapid actin reassembly for the initiation and retraction of protruded blebs. Live-cell imaging of membrane blebbing revealed that local reassembly of actin filaments occurred at Eps8- and activated ezrin-positive foci of membrane blebs. Furthermore, we found that a RhoA–ROCK–Rnd3 feedback loop determined the local reassembly sites of the actin cortex during membrane blebbing.Actin filaments usually lie beneath the plasma membrane. When the plasma membrane detaches from actin filaments, spherical protrusions of the membrane, termed blebs, form. Several lines of recent evidence suggest membrane blebs are used for cell migration under both physiological and pathological conditions. For example, migrating primordial germ cells (PGCs) use membrane blebs to migrate in zebrafish (1). Similarly, actively migrating PGCs exhibit membrane blebbing in Drosophila melanogaster embryos (2). Dictyostelium use membrane blebbing during chemotaxis (3, 4). Thus, membrane blebbing is widely used as a driving force of motility across species (4, 5). In addition, cancer cells use a membrane blebbing-associated mode of motility in metastasis (6). Cancer cells migrate without degrading the matrix by protruding membrane blebs in 3D extracellular matrixes (5, 7). Recently, cell physical confinement, down-regulation of cell adhesion to the extracellular matrix, and up-regulation of intrinsic cortical contraction forces were identified as key conditions for the induction of membrane blebbing-associated cell migration in many cell types including invasive cancer cells (8–10).However, the molecular mechanisms underlying membrane blebbing remain to be elucidated. Membrane blebs are initiated as a rapid protrusion of the plasma membrane, which is driven either by a change in intracellular hydrostatic pressure after local disruption of membrane–actin cortex interactions or by a local breakdown of actin filaments. Thereafter, actin filaments polymerize beneath the protruded membrane to halt bleb expansion. When actin filaments cover the protruded membrane, myosins are recruited to these filaments (11). It is unknown how actin cortex reassembly is triggered during these processes (11, 12), although there are at least two possibilities. One possibility is that actin filaments constitutively form beneath the plasma membrane and the plasma membrane stops extending when actin filaments are sufficiently reconstructed. The other possibility is that actin cortex reassembly is triggered by the activation of unidentified signaling when the cell senses a cytoskeleton-free plasma membrane region.In the present study, we addressed this issue using live-cell imaging and revealed that the rapid recovery of actin filaments at membrane blebs is promoted by epidermal growth factor receptor kinase substrate 8 (Eps8) and ezrin, and regulated by a RhoA–Rho-associated protein kinase (ROCK)–Rnd3 feedback loop.     

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.     

Spatiotemporal control of epithelial remodeling by regulated myosin phosphorylation

Spatiotemporally regulated actomyosin contractility generates the forces that drive epithelial cell rearrangements and tissue remodeling. Phosphorylation of the myosin II regulatory light chain (RLC) promotes the assembly of myosin monomers into active contractile filaments and is an essential mechanism regulating the level of myosin activity. However, the effects of phosphorylation on myosin localization, dynamics, and function during epithelial remodeling are not well understood. In Drosophila, planar polarized myosin contractility is required for oriented cell rearrangements during elongation of the body axis. We show that regulated myosin phosphorylation influences spatial and temporal properties of contractile behavior at molecular, cellular, and tissue length scales. Expression of myosin RLC variants that prevent or mimic phosphorylation both disrupt axis elongation, but have distinct effects at the molecular and cellular levels. Unphosphorylatable RLC produces fewer, slower cell rearrangements, whereas phosphomimetic RLC accelerates rearrangement and promotes higher-order cell interactions. Quantitative live imaging and biophysical approaches reveal that both phosphovariants reduce myosin planar polarity and mechanical anisotropy, altering the orientation of cell rearrangements during axis elongation. Moreover, the localized myosin activator Rho-kinase is required for spatially regulated myosin activity, even when the requirement for phosphorylation is bypassed by the expression of phosphomimetic myosin RLC. These results indicate that myosin phosphorylation influences both the level and the spatiotemporal regulation of myosin activity, linking molecular properties of myosin activity to tissue morphogenesis.Contractile assemblies of actin filaments and the nonmuscle myosin II motor protein produce mechanical forces that generate changes in cell shape during cytokinesis, cell movements during cell migration, and the dynamic remodeling of multicellular tissues (1–3). During development, spatiotemporal patterns of actomyosin contractility remodel simple epithelia into functional tissues with complex form and structure. Contractile force generation requires the assembly of inactive myosin monomers into active bipolar filaments (4). Phosphorylation of the myosin II regulatory light chain promotes myosin filament assembly and the movement of the myosin motor along actin filaments in vitro (5, 6) and is necessary for cytokinesis (7–9), oogenesis (7, 10, 11), and tissue morphogenesis (12–15). However, the in vivo effects of regulatory light chain phosphorylation on myosin localization, dynamics, and force generation, and how these properties are integrated to achieve complex changes in the shapes of tissues, are not well understood.Epithelial elongation in the Drosophila embryo is driven by cell rearrangements that are coordinated by spatiotemporal patterns of actomyosin contractility. Myosin contractility is spatially regulated in the plane of the epithelium (termed planar polarity), driving the planar polarized contraction of cell interfaces oriented close to perpendicular to the anterior–posterior axis (AP edges) (16–20). New interfaces preferentially form between dorsal and ventral cells (DV edges), resulting in oriented cell rearrangements that rapidly elongate the body axis of the animal from head to tail (16–18, 21). The myosin II activator Rho-associated protein kinase (Rho-kinase) localizes to regions of cells that display strong actomyosin activity during development (14, 15, 22) and promotes myosin contractility by phosphorylating the myosin regulatory light chain (RLC) and inactivating myosin phosphatase (23). Rho-kinase is required for localized myosin contractility during Drosophila axis elongation (14), neural tube closure in chick (22), and in other epithelia that undergo planar polarized cell behaviors (24, 25). Localized phosphorylation by Rho-kinase could therefore serve as an instructive cue that determines where and when myosin is activated within the cell. However, constitutively active myosin variants that evade this upstream regulation are sufficient to support normal myosin function during cytokinesis (26, 27), oogenesis (7), and wing hair formation (12), indicating that in some contexts regulated myosin phosphorylation is dispensable for morphogenesis. This raises the alternative possibility that the primary requirement for phosphorylation is to activate myosin, and additional mechanisms regulate the spatiotemporal patterns of myosin localization within epithelia.Here we use time-lapse imaging and biophysical approaches to analyze the role of myosin phosphorylation in polarized cell rearrangements during Drosophila axis elongation. Myosin II regulatory light chain variants that prevent or mimic phosphorylation both disrupt axis elongation, but have distinct effects on myosin localization, dynamics, and cell behavior. These results indicate that regulated myosin phosphorylation provides an instructive cue that directs both the magnitude and spatiotemporal pattern of myosin activity during epithelial remodeling, linking molecular-scale myosin activity to cell behavior and tissue morphogenesis.     

Predictive Value of Brain Natriuretic Peptides in Patients on Peritoneal Dialysis: Results from the ADEMEX Trial

Background and objectives: Natriuretic peptides have been suggested to be of value in risk stratification in dialysis patients. Data in patients on peritoneal dialysis remain limited.Design, setting, participants, & measurements: Patients of the ADEMEX trial (ADEquacy of peritoneal dialysis in MEXico) were randomized to a control group [standard 4 × 2L continuous ambulatory peritoneal dialysis (CAPD); n = 484] and an intervention group (CAPD with a target creatinine clearance ≥60L/wk/1.73 m²; n = 481). Natriuretic peptides were measured at baseline and correlated with other parameters as well as evaluated for effects on patient outcomes.Results: Control group and intervention group were comparable at baseline with respect to all measured parameters. Baseline values of natriuretic peptides were elevated and correlated significantly with levels of residual renal function but not with body size or diabetes. Baseline values of N-terminal fragment of B-type natriuretic peptide (NT-proBNP) but not proANP(1–30), proANP(31–67), or proANP(1–98) were independently highly predictive of overall survival and cardiovascular mortality. Volume removal was also significantly correlated with patient survival.Conclusions. NT-proBNP have a significant predictive value for survival of CAPD patients and may be of value in guiding risk stratification and potentially targeted therapeutic interventions.Plasma levels of cardiac natriuretic peptides are elevated in patients with chronic kidney disease, owing to impairment of renal function, hypertension, hypervolemia, and/or concomitant heart disease (1–7). Atrial natriuretic peptide (ANP) and particularly brain natriuretic peptide (BNP) levels are linked independently to left ventricular mass (3–5,8–16) and function (3,6–17) and predict total and cardiovascular mortality (1,3,8,10,12,18) as well as cardiac events (12,19). ANP and BNP decrease significantly during hemodialysis treatment but increase again during the interdialytic interval (1,2,4,6,7,14,17,20–23). Levels in patients on peritoneal dialysis (PD) have been found to be lower than in patients on hemodialysis (11,24–26), but the correlations with left ventricular function and structure are maintained in both types of dialysis modalities (11,15,27,28).The high mortality of patients on peritoneal dialysis and the failure of dialytic interventions to alter this mortality (29,30) necessitate renewed attention into novel methods of stratification and identification of patients at highest risk to be targeted for specific interventions. Cardiac natriuretic peptides are increasingly considered to fulfill this role in nonrenal patients. Evaluations of cardiac natriuretic peptides in patients on PD have been limited by small numbers (3,9,11,12,15,24–26) and only one study examined correlations between natriuretic peptide levels and outcomes (12). The PD population enrolled in the ADEMEX trial offered us the opportunity to evaluate cardiac natriuretic peptides and their value in predicting outcomes in the largest clinical trial ever performed on PD (29,30). It is hoped that such an evaluation would identify patients at risk even in the absence of overt clinical disease and hence facilitate or encourage interventions with salutary outcomes.     

Structural basis for membrane recruitment and allosteric activation of cytohesin family Arf GTPase exchange factors

Membrane recruitment of cytohesin family Arf guanine nucleotide exchange factors depends on interactions with phosphoinositides and active Arf GTPases that, in turn, relieve autoinhibition of the catalytic Sec7 domain through an unknown structural mechanism. Here, we show that Arf6-GTP relieves autoinhibition by binding to an allosteric site that includes the autoinhibitory elements in addition to the PH domain. The crystal structure of a cytohesin-3 construct encompassing the allosteric site in complex with the head group of phosphatidyl inositol 3,4,5-trisphosphate and N-terminally truncated Arf6-GTP reveals a large conformational rearrangement, whereby autoinhibition can be relieved by competitive sequestration of the autoinhibitory elements in grooves at the Arf6/PH domain interface. Disposition of the known membrane targeting determinants on a common surface is compatible with multivalent membrane docking and subsequent activation of Arf substrates, suggesting a plausible model through which membrane recruitment and allosteric activation could be structurally integrated.Guanine nucleotide exchange factors (GEFs) activate GTPases by catalyzing exchange of GDP for GTP (1). Because many GEFs are recruited to membranes through interactions with phospholipids, active GTPases, or other membrane-associated proteins (1–5), GTPase activation can be restricted or amplified by spatial–temporal overlap of GEFs with binding partners. GEF activity can also be controlled by autoregulatory mechanisms, which may depend on membrane recruitment (6–11). Structural relationships between these mechanisms are poorly understood.Arf GTPases function in trafficking and cytoskeletal dynamics (5, 12, 13). Membrane partitioning of a myristoylated (myr) N-terminal amphipathic helix primes Arfs for activation by Sec7 domain GEFs (14–17). Cytohesins comprise a metazoan Arf GEF family that includes the mammalian proteins cytohesin-1 (Cyth1), ARNO (Cyth2), and Grp1 (Cyth3). The Drosophila homolog steppke functions in insulin-like growth factor signaling, whereas Cyth1 and Grp1 have been implicated in insulin signaling and Glut4 trafficking, respectively (18–20). Cytohesins share a modular architecture consisting of heptad repeats, a Sec7 domain with exchange activity for Arf1 and Arf6, a PH domain that binds phosphatidyl inositol (PI) polyphosphates, and a C-terminal helix (CtH) that overlaps with a polybasic region (PBR) (21–28). The overlapping CtH and PBR will be referred to as the CtH/PBR. The phosphoinositide specificity of the PH domain is influenced by alternative splicing, which generates diglycine (2G) and triglycine (3G) variants differing by insertion of a glycine residue in the β1/β2 loop (29). Despite similar PI(4,5)P₂ (PIP₂) affinities, the 2G variant has 30-fold higher affinity for PI(3,4,5)P₃ (PIP₃) (30). In both cases, PIP₃ is required for plasma membrane (PM) recruitment (23, 26, 31–33), which is promoted by expression of constitutively active Arf6 or Arl4d and impaired by PH domain mutations that disrupt PIP₃ or Arf6 binding, or by CtH/PBR mutations (8, 34–36).Cytohesins are autoinhibited by the Sec7-PH linker and CtH/PBR, which obstruct substrate binding (8). Autoinhibition can be relieved by Arf6-GTP binding in the presence of the PIP₃ head group (8). Active myr-Arf1 and myr-Arf6 also stimulate exchange activity on PIP₂-containing liposomes (37). Whether this effect is due to relief of autoinhibition per se or enhanced membrane recruitment is not yet clear. Phosphoinositide recognition by PH domains, catalysis of nucleotide exchange by Sec7 domains, and autoinhibition in cytohesins are well characterized (8, 16, 17, 30, 38–43). How Arf-GTP binding relieves autoinhibition and promotes membrane recruitment is unknown. Here, we determine the structural basis for relief of autoinhibition and investigate potential mechanistic relationships between allosteric regulation, phosphoinositide binding, and membrane targeting.     

Differential gradients of interaction affinities drive efficient targeting and recycling in the GET pathway

Efficient and accurate localization of membrane proteins requires a complex cascade of interactions between protein machineries. This requirement is exemplified in the guided entry of tail-anchored (TA) protein (GET) pathway, where the central targeting factor Get3 must sequentially interact with three distinct binding partners to ensure the delivery of TA proteins to the endoplasmic reticulum (ER) membrane. To understand the molecular principles that provide the vectorial driving force of these interactions, we developed quantitative fluorescence assays to monitor Get3–effector interactions at each stage of targeting. We show that nucleotide and substrate generate differential gradients of interaction energies that drive the ordered interaction of Get3 with successive effectors. These data also provide more molecular details on how the targeting complex is captured and disassembled by the ER receptor and reveal a previously unidentified role for Get4/5 in recycling Get3 from the ER membrane at the end of the targeting reaction. These results provide general insights into how complex protein interaction cascades are coupled to energy inputs in biological systems.Membrane proteins comprise ∼30% of the proteome; their efficient and accurate localization is crucial for the structure and function of all cells. Although the well-studied cotranslational signal recognition particle pathway delivers numerous endoplasmic reticulum (ER) -destined proteins (1), many membrane proteins use posttranslational targeting pathways with mechanisms that are far less well understood. A salient example is tail-anchored (TA) proteins, which comprise 3–5% of the eukaryotic membrane proteome and play essential roles in numerous processes, including protein translocation, vesicular trafficking, quality control, and apoptosis (2–5). Because their sole transmembrane domain is at the extreme C terminus, TA proteins cannot engage the cotranslational signal recognition particle machinery and instead, must use posttranslational pathways for localization (6).In the guided entry of TA protein (GET) pathway, TA proteins are initially captured by the yeast cochaperone Sgt2 (or mammalian SGTA) (2, 7). The Get4/5 complex then enables loading of the TA substrate from Sgt2 onto Get3 (or mammalian TRC40), the central targeting factor (7–9). The Get3/TA complex binds a receptor complex on the ER membrane comprised of Get1 and Get2, through which the TA protein is released from Get3 and inserted into the membrane (10–12). Dissociation from Get1/2 is then needed to recycle Get3 for additional rounds of targeting (11–13). Knockout of Get3 (or TRC40) confers stress sensitivity in yeast and embryonic lethality in mammals, underscoring its essential role in the proper functioning of the cell (10, 14, 15).TA protein targeting is driven by the ATPase cycle of Get3, a member of the signal recognition particle, MinD and BioD class of nucleotide hydrolases (8, 16). Crystallographic studies revealed that Get3 is an obligate homodimer, in which the ATPase domains bridge the dimer interface and are connected to helical domains (17, 18). Notably, the conformation of Get3 can be tuned by its nucleotide state, the TA substrate, and its binding partners (11, 12, 17, 19). Apo-Get3 is in an open conformation, in which the helical domains are disconnected (18). ATP biases Get3 to more closed structures, in which the helical domains form a contiguous hydrophobic surface implicated in TA protein binding (17, 18, 20). The Get4/5 complex further locks Get3 into an occluded conformation, in which ATP is tightly bound, but its hydrolysis is delayed, priming Get3 into the optimal state to capture the TA substrate (19, 21). TA proteins induce additional association of Get3 dimers to form a closed tetramer, which stimulates rapid ATP hydrolysis and delays ADP release (19, 22). Finally, Get1 strongly binds apo-Get3 in the open conformation (see below), likely at the end of the targeting reaction (11, 12, 23).The GET pathway demands a sequential cascade of interactions of Get3 with three distinct binding partners: the Get4 subunit in the Get4/5 complex and the Get1 and Get2 subunits in the Get1/2 receptor complex. All three partners share overlapping binding sites on Get3 (Fig. S1) (21), raising intriguing questions as to the mechanisms that ensure the high spatial and temporal accuracy of these protein interactions. For example, Get3 must first interact with Get4/5 in the cytosol to facilitate the loading of TA substrate (7, 9). It is unclear what then drives the release of Get3 from Get4/5 and enables its transit to the ER membrane, where it interacts with the Get1/2 receptor instead.Similarly, how Get3 and the Get3/TA complex transit between different subunits of the Get1/2 receptor at the ER membrane remains unclear. Get1/2 (WRB/CAML in mammals) is necessary and sufficient for TA protein insertion at the ER membrane (12, 13, 24, 25). Crystallographic analyses revealed that Get1 binds strongly to apo- or ADP-bound Get3 in the open conformation (11, 12, 23), whereas Get2 can bind Get3 in semiclosed or closed states (11, 12). In vitro reconstitution experiments showed that high concentrations of Get1 but not Get2 can trigger substrate release from Get3 (12). These observations led to the model that Get2 first captures Get3, whereas Get1 is responsible for disassembling the targeting complex (2, 13). Nevertheless, the subunit that is responsible for capturing the Get3/TA targeting complex has not been experimentally addressed, and whether Get1 or Get2 can discriminate different substrate-bound states of Get3 also has not been addressed.At the end of targeting, Get1 is tightly bound to apo-Get3 (11–13). Experiments with the cytosolic domain (CD) of Get1 show that its interaction with Get3 is strongly antagonized by ATP, leading to the current model that ATP drives the recycling of Get3 from the ER membrane (11, 12). However, two observations raise difficulties with this minimal model. In experiments with intact ER membranes or Get1/2 proteoliposomes (PLs), ATP is insufficient to completely release Get3 from the membrane (12, 13). Furthermore, the tight interaction of Get1 with Get3 raises the possibility that their dissociation is slow (11), which could pose potential barriers for subsequent rounds of TA protein targeting.To address these issues, we developed fluorescence assays to report on the interaction of Get3 with its effectors. Quantitative measurements show that both substrate and nucleotide regulate the interaction of Get3 with Get4/5 and Get1/2, generating differential gradients of interaction energies that drive the ordered transit of Get3 from one binding partner to the next. These results also reveal an active role of ATP in displacing Get3 from Get1, which together with Get4/5, ensures the effective recycling of Get3 back to the cytosol.     

N terminus of ASPP2 binds to Ras and enhances Ras/Raf/MEK/ERK activation to promote oncogene-induced senescence

The ASPP2 (also known as 53BP2L) tumor suppressor is a proapoptotic member of a family of p53 binding proteins that functions in part by enhancing p53-dependent apoptosis via its C-terminal p53-binding domain. Mounting evidence also suggests that ASPP2 harbors important nonapoptotic p53-independent functions. Structural studies identify a small G protein Ras-association domain in the ASPP2 N terminus. Because Ras-induced senescence is a barrier to tumor formation in normal cells, we investigated whether ASPP2 could bind Ras and stimulate the protein kinase Raf/MEK/ERK signaling cascade. We now show that ASPP2 binds to Ras–GTP at the plasma membrane and stimulates Ras-induced signaling and pERK1/2 levels via promoting Ras–GTP loading, B-Raf/C-Raf dimerization, and C-Raf phosphorylation. These functions require the ASPP2 N terminus because BBP (also known as 53BP2S), an alternatively spliced ASPP2 isoform lacking the N terminus, was defective in binding Ras–GTP and stimulating Raf/MEK/ERK signaling. Decreased ASPP2 levels attenuated H-RasV12–induced senescence in normal human fibroblasts and neonatal human epidermal keratinocytes. Together, our results reveal a mechanism for ASPP2 tumor suppressor function via direct interaction with Ras–GTP to stimulate Ras-induced senescence in nontransformed human cells.ASPP2, also known as 53BP2L, is a tumor suppressor whose expression is altered in human cancers (1). Importantly, targeting of the ASPP2 allele in two different mouse models reveals that ASPP2 heterozygous mice are prone to spontaneous and γ-irradiation–induced tumors, which rigorously demonstrates the role of ASPP2 as a tumor suppressor (2, 3). ASPP2 binds p53 via the C-terminal ankyrin-repeat and SH3 domain (4–6), is damage-inducible, and can enhance damage-induced apoptosis in part through a p53-mediated pathway (1, 2, 7–10). However, it remains unclear what biologic pathways and mechanisms mediate ASPP2 tumor suppressor function (1). Indeed, accumulating evidence demonstrates that ASPP2 also mediates nonapoptotic p53-independent pathways (1, 3, 11–15).The induction of cellular senescence forms an important barrier to tumorigenesis in vivo (16–21). It is well known that oncogenic Ras signaling induces senescence in normal nontransformed cells to prevent tumor initiation and maintain complex growth arrest pathways (16, 18, 21–24). The level of oncogenic Ras activation influences its capacity to activate senescence; high levels of oncogenic H-RasV12 signaling leads to low grade tumors with senescence markers, which progress to invasive cancers upon senescence inactivation (25). Thus, tight control of Ras signaling is critical to ensure the proper biologic outcome in the correct cellular context (26–28).The ASPP2 C terminus is important for promoting p53-dependent apoptosis (7). The ASPP2 N terminus may also suppress cell growth (1, 7, 29–33). Alternative splicing can generate the ASPP2 N-terminal truncated protein BBP (also known as 53BP2S) that is less potent in suppressing cell growth (7, 34, 35). Although the ASPP2 C terminus mediates nuclear localization, full-length ASPP2 also localizes to the cytoplasm and plasma membrane to mediate extranuclear functions (7, 11, 12, 36). Structural studies of the ASPP2 N terminus reveal a β–Grasp ubiquitin-like fold as well as a potential Ras-binding (RB)/Ras-association (RA) domain (32). Moreover, ASPP2 can promote H-RasV12–induced senescence (13, 15). However, the molecular mechanism(s) of how ASPP2 directly promotes Ras signaling are complex and remain to be completely elucidated.Here, we explore the molecular mechanisms of how Ras-signaling is enhanced by ASPP2. We demonstrate that ASPP2: (i) binds Ras-GTP and stimulates Ras-induced ERK signaling via its N-terminal domain at the plasma membrane; (ii) enhances Ras-GTP loading and B-Raf/C-Raf dimerization and forms a ASPP2/Raf complex; (iii) stimulates Ras-induced C-Raf phosphorylation and activation; and (iv) potentiates H-RasV12–induced senescence in both primary human fibroblasts and neonatal human epidermal keratinocytes. These data provide mechanistic insight into ASPP2 function(s) and opens important avenues for investigation into its role as a tumor suppressor in human cancer.