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.     

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.     

Myosin-binding protein C displaces tropomyosin to activate cardiac thin filaments and governs their speed by an independent mechanism

Myosin-binding protein C (MyBP-C) is an accessory protein of striated muscle thick filaments and a modulator of cardiac muscle contraction. Defects in the cardiac isoform, cMyBP-C, cause heart disease. cMyBP-C includes 11 Ig- and fibronectin-like domains and a cMyBP-C-specific motif. In vitro studies show that in addition to binding to the thick filament via its C-terminal region, cMyBP-C can also interact with actin via its N-terminal domains, modulating thin filament motility. Structural observations of F-actin decorated with N-terminal fragments of cMyBP-C suggest that cMyBP-C binds to actin close to the low Ca²⁺ binding site of tropomyosin. This suggests that cMyBP-C might modulate thin filament activity by interfering with tropomyosin regulatory movements on actin. To determine directly whether cMyBP-C binding affects tropomyosin position, we have used electron microscopy and in vitro motility assays to study the structural and functional effects of N-terminal fragments binding to thin filaments. 3D reconstructions suggest that under low Ca²⁺ conditions, cMyBP-C displaces tropomyosin toward its high Ca²⁺ position, and that this movement corresponds to thin filament activation in the motility assay. At high Ca²⁺, cMyBP-C had little effect on tropomyosin position and caused slowing of thin filament sliding. Unexpectedly, a shorter N-terminal fragment did not displace tropomyosin or activate the thin filament at low Ca²⁺ but slowed thin filament sliding as much as the larger fragments. These results suggest that cMyBP-C may both modulate thin filament activity, by physically displacing tropomyosin from its low Ca²⁺ position on actin, and govern contractile speed by an independent molecular mechanism.Myosin-binding protein C (MyBP-C) is an accessory protein of vertebrate striated muscle thick filaments (1) that is known to modulate cardiac muscle contraction (2). The skeletal isoform includes 10 Ig-like (Ig) and fibronectin type 3-like (Fn) domains, numbered C1 through C10 from the N terminus, together with a MyBP-C-specific motif (the M-domain) between C1 and C2 and a Pro-Ala-rich sequence at the N terminus. The cardiac isoform (cMyBP-C) has an additional N-terminal Ig domain (C0), four phosphorylation sites in the M-domain, and a 28-residue insert in the C5 domain (3) (Fig. 1). MyBP-C binds to the thick filament in the C-zone of the sarcomeric A-band (4) via its C-terminal domains (C8–C10) (5), whereas its N-terminal region contains binding sites for myosin S2 (6–10) and the myosin regulatory light chain (11).Open in a separate windowFig. 1.Schematic of cMyBP-C and the expressed N-terminal fragments C0C3, C0C2, and C0C1f used in this study. cMyBP-C consists of 8 Ig and 3 Fn domains together with a cMyBP-C-specific M-domain, containing a phosphorylation region (orange) with 4 phosphorylatable serines (P), a ProAla-rich domain, and a cardiac-specific insert (blue) in the C5 domain.In addition to binding to myosin, MyBP-C also interacts with actin (12) and with thin filaments (13) via its N-terminal region (9, 10, 14–17; cf. 18). In the in vitro motility assay, actin filament sliding over myosin is slowed by N-terminal fragments of cMyBP-C to the same extent as whole cMyBP-C (19), possibly by slowing the myosin detachment rate from actin (19) or tethering the thick to the thin filament (16, 19). In an assay closer to the in vivo situation, the sliding of F-actin over native cardiac thick filaments was slowed specifically in the C-zone, and this slowing was ablated by removal of C0C1 and the first 17 amino acids of the M-domain [known as C0C1f (16); Fig. 1] (20). On the basis of yeast 2 hybrid experiments, it was concluded that the C1 and M domains were necessary for actin binding and that replacement of endogenous cMyBP-C with actin binding-ablated cMyBP-C resulted in its abnormal sarcomeric distribution and disturbance of the sarcomeric structure (9). These in vitro demonstrations of actin binding are supported by electron tomographic observations showing MyBP-C extending from the thick to the thin filaments in the intact sarcomere, consistent with a model in which the N terminus of MyBP-C binds to the thin filament (21, 22). Together, these results suggest that actin binding is physiologically relevant and that the slowing of actin filament sliding is one possible mechanism by which cMyBP-C modulates cardiac contractility (5, 23, 24).The structural basis of cMyBP-C’s N-terminal binding to F-actin has been studied in several ways. Neutron scattering and NMR titration analysis of F-actin decorated with the N-terminal fragment C0C2 (Fig. 1) suggested that C0C2 binds to subdomain 1 (SD1) and the DNase loop of actin (25) via key regions within C0 and C1 (10). More direct observations by negative stain electron microscopy (26) and 3D reconstruction (27) suggest that C0 and C1 bind to SD1 of actin, whereas the M domain crosses over SD2, possibly binding to the next SD1, and C2 and C3 lie above the surface of the filament (27, 28).The position of cMyBP-C binding on actin SD1 suggests that in addition to inhibiting actomyosin interactions, it might also affect thin filament regulation by interfering with the binding of tropomyosin/troponin (Tm/Tn) to actin. Tm and Tn regulate muscle contraction by movement of Tm in response to calcium binding by Tn (29). At low Ca²⁺, Tm lies on SD1, where it sterically blocks the binding of myosin to actin (the “blocked” position); the consequent inhibition of actin-myosin interaction leads to muscle relaxation. On activation, Tn binds Ca²⁺, causing Tm to move onto actin SD3 (the “closed” position), exposing myosin binding sites on SD1 and initiating crossbridge cycling and contraction (29–35). When a model of Tm in its low Ca²⁺ (blocked) state is positioned on the reconstruction of F-actin decorated with C0C3, it appears to clash with cMyBP-C’s C0 and C1 domains, suggesting that cMyBP-C and Tm might compete for binding to SD1 in the relaxed thin filament (25, 27, 28). As a consequence, cMyBP-C might be expected to activate the thin filament by physically preventing Tm from assuming its blocked position (25, 27, 28). Motility (19) and solution kinetics studies (36, 37) support this concept, showing that the N-terminal C1C2 fragment activates thin filaments in low Ca²⁺ similarly to rigor heads.Here we have investigated cMyBP-C’s potential to modulate cardiac contractility by contrasting mechanisms; that is, activating the thin filament and inhibiting maximal actomyosin mechanical activity. First we used negative-staining EM and 3D reconstruction to investigate whether cMyBP-C displaces Tm at low Ca²⁺, by decorating regulated thin filaments (containing F-actin, Tm and Tn) with C0C2. In parallel experiments, we determined the functional consequences of such N-terminal fragments on regulated thin filament activation and sliding velocities in an in vitro motility assay. We find clear structural evidence for displacement of Tm toward the high Ca²⁺ (closed) position when C0C2 binds to thin filaments under low Ca²⁺ conditions, suggesting that N-terminal binding should activate the thin filament; this was confirmed in the motility assay. We also demonstrate that C0C2 has no effect on Tm position under high Ca²⁺ conditions but inhibits maximal sliding velocity in the motility assay. Interestingly, a smaller fragment (C0C1f; Fig. 1) binds to the thin filament under low Ca²⁺ conditions but does not displace Tm or activate thin filament sliding. However, C0C1f still inhibits thin filament sliding at high Ca²⁺ to the same extent as the larger N-terminal fragment. These results suggest that cMyBP-C may play two physiological roles in intact muscle, displacing Tm from the “blocked” position at low Ca²⁺ to modulate thin filament activation, and governing maximal sliding velocity at high Ca²⁺ by a potentially independent molecular mechanism.     

Myosin binding protein-C activates thin filaments and inhibits thick filaments in heart muscle cells

Myosin binding protein-C (MyBP-C) is a key regulatory protein in heart muscle, and mutations in the MYBPC3 gene are frequently associated with cardiomyopathy. However, the mechanism of action of MyBP-C remains poorly understood, and both activating and inhibitory effects of MyBP-C on contractility have been reported. To clarify the function of the regulatory N-terminal domains of MyBP-C, we determined their effects on the structure of thick (myosin-containing) and thin (actin-containing) filaments in intact sarcomeres of heart muscle. We used fluorescent probes on troponin C in the thin filaments and on myosin regulatory light chain in the thick filaments to monitor structural changes associated with activation of demembranated trabeculae from rat ventricle by the C1mC2 region of rat MyBP-C. C1mC2 induced larger structural changes in thin filaments than calcium activation, and these were still present when active force was blocked with blebbistatin, showing that C1mC2 directly activates the thin filaments. In contrast, structural changes in thick filaments induced by C1mC2 were smaller than those associated with calcium activation and were abolished or reversed by blebbistatin. Low concentrations of C1mC2 did not affect resting force but increased calcium sensitivity and reduced cooperativity of force and structural changes in both thin and thick filaments. These results show that the N-terminal region of MyBP-C stabilizes the ON state of thin filaments and the OFF state of thick filaments and lead to a novel hypothesis for the physiological role of MyBP-C in the regulation of cardiac contractility.Muscle contraction is driven by the relative sliding of the actin-containing thin filaments along the myosin-containing thick filaments arranged in a parallel array in the muscle sarcomere (Fig. 1A). Filament sliding in turn is driven by a structural change in the myosin head domains (Fig. 1B) while they are bound to actin, coupled to the hydrolysis of ATP (1). Contraction of skeletal and cardiac muscle is triggered by calcium binding to troponin in the thin filaments, accompanied by a change in the structure of the thin filaments that permits myosin head binding (2). However, the strength and dynamics of contraction are modulated by posttranslational modifications in other sarcomeric proteins, including the myosin regulatory light chain (RLC) (3), which is part of the myosin head, and myosin binding protein-C (4–6) (MyBP-C) (Fig. 1B). In an emerging concept of thick filament regulation in striated muscle that is analogous to myosin-linked regulation in smooth muscle (7–11), RLC and MyBP-C are thought to modulate contraction by controlling the conformation of the myosin heads.Open in a separate windowFig. 1.Sarcomere location and domain architecture of MyBP-C. (A) C-zone (green) of the thick filament in relation to its proximal (P) and distal (D) regions and the thin filament (gray). (B) Cartoon representation of MyBP-C (green) anchored to the thick filament backbone (purple) via its C-terminal domains; myosin heads are pink and troponin is yellow. (C) Domain organization and interactions of MyBP-C.According to this concept, the thick filament has an OFF state in which the myosin heads are folded back against its surface (Fig. 1B), rendering them unavailable for interaction with actin, and an ON state in which the heads are released from the thick filament surface and made available for actin binding. The physiological and pathological significance of thick filament regulation and its relationship to the well-studied thin filament mechanisms remain poorly understood, but much recent attention has focused on MyBP-C for two main reasons. First, mutations in the cardiac MYBPC3 gene are commonly associated with hypertrophic cardiomyopathy (12, 13), and this association has driven a wide range of studies at the molecular, cellular, and whole-animal levels aimed at understanding the etiology of MYBPC3-linked disease. Second, although MyBP-C is a constitutive component of the thick filament, there is a large body of evidence that it can also bind the thin filaments (14, 15), raising the possibility that one role of MyBP-C may be to synchronize the regulatory states of the thin and thick filaments (11, 15–17).MyBP-C is localized to the central region or “C-zone” of each half-thick filament (Fig. 1A), appearing in nine transverse stripes with a 43-nm periodicity closely matching that of the myosin heads (Fig. 1B) (10). MyBP-C has 11 Ig-like or fibronectin-like domains (Fig. 1C) denoted C0–C10, with additional linking sequences, notably the MyBP-C “motif” or “m” domain between C1 and C2 and the proline/alanine-rich (P/A) linker between C0 and C1. The m domain has multiple phosphorylation sites (4–6). Constitutive binding to the thick filament is mediated by interactions of domains C8–C10 with myosin and titin. The C1mC2 region binds to the coiled-coil subfragment-2 (S2) domain of myosin adjacent to the myosin heads, and this interaction is abolished by MyBP-C phosphorylation (5); the C0 domain binds to the RLC in the myosin head itself (18). The N-terminal domains of MyBP-C also bind to actin in a phosphorylation-dependent manner (14, 15) (Fig. 1B), and EM and X-ray studies on intact sarcomeres of skeletal muscle suggest that MyBP-C binds to thin filaments under relaxing conditions (10, 11).The function of MyBP-C and the mechanisms underlying its modulation in cardiomyopathy remain poorly understood, however. Ablation of MyBP-C in a knockout mouse model leads to a hypertrophic phenotype associated with impaired contractile function (19), but cardiomyocytes isolated from these mice exhibit increased power output during working contractions (20). A range of studies at the isolated protein and cellular levels have led to the concept that MyBP-C exerts a predominantly inhibitory effect on contractility mediated through two distinct mechanisms (15, 16, 21). MyBP-C may tether myosin heads to the surface of the thick filament, preventing their interaction with actin, and its N terminus may bind to thin filaments, inhibiting interfilament sliding at low load. Other studies, however, have demonstrated an activating effect of MyBP-C mediated by binding of its N-terminal domains to the thin filament. N-terminal fragments of MyBP-C enhance force production in skinned cardiac muscle cells and motility in isolated filament preparations at zero or submaximal calcium concentrations (22–25). The same effect is observed in cardiomyocytes from MyBP-C knockout mice (22), suggesting that the activating effect is not due to competitive removal of an inhibitory effect of native MyBP-C.To resolve these apparently contradictory hypotheses about the physiological function of the N-terminal domains of MyBP-C, we determined the structural changes in the thick and thin filaments of intact sarcomeres in heart muscle cells induced by N-terminal MyBP-C fragments using bifunctional rhodamine probes on RLC and troponin C (TnC) (26). These probes allowed the structural changes in both types of filament to be directly compared with those associated with calcium activation and myosin head binding in the native environment of the cardiac muscle sarcomere. The results lead to a model for the physiological function of MyBP-C that integrates the regulatory roles of the thin and thick filaments and the inhibitory and activating effects of MyBP-C at the level of the intact sarcomere.     

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.     

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.     

Self-assembly of amphiphilic Janus dendrimers into uniform onion-like dendrimersomes with predictable size and number of bilayers

A constitutional isomeric library synthesized by a modular approach has been used to discover six amphiphilic Janus dendrimer primary structures, which self-assemble into uniform onion-like vesicles with predictable dimensions and number of internal bilayers. These vesicles, denoted onion-like dendrimersomes, are assembled by simple injection of a solution of Janus dendrimer in a water-miscible solvent into water or buffer. These dendrimersomes provide mimics of double-bilayer and multibilayer biological membranes with dimensions and number of bilayers predicted by the Janus compound concentration in water. The simple injection method of preparation is accessible without any special equipment, generating uniform vesicles, and thus provides a promising tool for fundamental studies as well as technological applications in nanomedicine and other fields.Most living organisms contain single-bilayer membranes composed of lipids, glycolipids, cholesterol, transmembrane proteins, and glycoproteins (1). Gram-negative bacteria (2, 3) and the cell nucleus (4), however, exhibit a strikingly special envelope that consists of a concentric double-bilayer membrane. More complex membranes are also encountered in cells and their various organelles, such as multivesicular structures of eukaryotic cells (5) and endosomes (6), and multibilayer structures of endoplasmic reticulum (7, 8), myelin (9, 10), and multilamellar bodies (11, 12). This diversity of biological membranes inspired corresponding biological mimics. Liposomes (Fig. 1) self-assembled from phospholipids are the first mimics of single-bilayer biological membranes (13–16), but they are polydisperse, unstable, and permeable (14). Stealth liposomes coassembled from phospholipids, cholesterol, and phospholipids conjugated with poly(ethylene glycol) exhibit improved stability, permeability, and mechanical properties (17–20). Polymersomes (21–24) assembled from amphiphilic block copolymers exhibit better mechanical properties and permeability, but are not always biocompatible and are polydisperse. Dendrimersomes (25–28) self-assembled from amphiphilic Janus dendrimers and minidendrimers (26–28) have also been elaborated to mimic single-bilayer biological membranes. Amphiphilic Janus dendrimers take advantage of multivalency both in their hydrophobic and hydrophilic parts (23, 29–32). Dendrimersomes are assembled by simple injection (33) of a solution of an amphiphilic Janus dendrimer (26) in a water-soluble solvent into water or buffer and produce uniform (34), impermeable, and stable vesicles with excellent mechanical properties. In addition, their size and properties can be predicted by their primary structure (27). Amphiphilic Janus glycodendrimers self-assemble into glycodendrimersomes that mimic the glycan ligands of biological membranes (35). They have been demonstrated to be bioactive toward biomedically relevant bacterial, plant, and human lectins, and could have numerous applications in nanomedicine (20).Open in a separate windowFig. 1.Strategies for the preparation of single-bilayer vesicles and multibilayer onion-like vesicles.More complex and functional cell mimics such as multivesicular vesicles (36, 37) and multibilayer onion-like vesicles (38–40) have also been discovered. Multivesicular vesicles compartmentalize a larger vesicle (37) whereas multibilayer onion-like vesicles consist of concentric alternating bilayers (40). Currently multibilayer vesicles are obtained by very complex and time-consuming methods that do not control their size (39) and size distribution (40) in a precise way. Here we report the discovery of “single–single” (28) amphiphilic Janus dendrimer primary structures that self-assemble into uniform multibilayer onion-like dendrimersomes (Fig. 1) with predictable size and number of bilayers by simple injection of their solution into water or buffer.     

Solid-phase peptide synthesis and solid-phase fragment coupling mediated by isonitriles

The synthesis of polypeptides on solid phase via mediation by isonitriles is described. The acyl donor is a thioacid, which presumably reacts with the isonitrile to generate a thio-formimidate carboxylate mixed anhydride intermediate. Applications of this chemistry to reiterative solid-phase peptide synthesis as well as solid-phase fragment coupling are described.Amide bond formations are arguably among the most important constructions in organic chemistry (1, 2). The centrality of the amide linkage, as found in polypeptides and proteins, in the maintenance of life hardly needs restatement. Numerous strategies, resulting in a vast array of protocols to synthesize biologically active polypeptides and proteins, have been demonstrated (3, 4). Central to reiterative polypeptide bond formations was the discovery and remarkable development of solid-phase peptide synthesis (SPPS) (5, 6). The extraordinary impact of SPPS in fostering enhanced access to homogeneous polypeptides is clear to everyone in the field.As we have described elsewhere, by classical, mechanistic reasoning, we were led to conjecture about some hitherto-unexplored possibilities relevant to the chemistry of isonitriles (7–14). It was anticipated that isonitriles might be able to mediate the acylation of amines, thus giving rise to amides (15). Early experiments focused on free carboxylic acids as the acylating agents. As our studies progressed, it was found that the combination of thioacids, amines, and isonitriles leads to the efficient formation of amide bonds under stoichiometric or near-stoichiometric conditions (7–13, 16, 17). Although there remain unresolved issues of detail and nuance, the governing mechanism for amide formation under these conditions involves reaction of the thioacid, 1, with an isonitrile, 2, to generate a thio-formimidate carboxylate mixed anhydride (thio-FCMA), 3, which is intercepted by the “acyl-accepting” amine to generate amide, 5, and thioformamide, 6 (Fig. 1). The efficiency of the amidation was further improved through the use of hydroxybenzotriazole (HOBt) (18), which could well give rise to HOBt ester 7, although this pathway has not been mechanistically proven.Open in a separate windowFig. 1.Isonitrile-mediated amidation; structure of OT.The potentialities of the isonitrile-mediated amidation method were foreshadowed via its application to the synthesis of cyclosporine (19). The power of the method was particularly well demonstrated in the context of our recent total synthesis of oxytocin (OT) (20), wherein isonitrile mediation was used in each of the peptide bond constructions, leading to the synthesis of the hormone in high yield and excellent purity. This nonapeptide is involved in a range of biological functions including parturition and lactation (21, 22). Signaling of OT to its receptor (OTR) is apparently an important factor in quality maintenance of various CNS functions (23). The ability to synthesize such modestly sized, but bio-impactful peptides in both native (wild-type) form, and as strategically modified variants, is one of the current missions of our laboratory, with the objective of possible applications to the very serious problem of autism (24–26).     

Allosteric N-WASP activation by an inter-SH3 domain linker in Nck

Actin filament networks assemble on cellular membranes in response to signals that locally activate neural Wiskott–Aldrich-syndrome protein (N-WASP) and the Arp2/3 complex. An inactive conformation of N-WASP is stabilized by intramolecular contacts between the GTPase binding domain (GBD) and the C helix of the verprolin-homology, connector-helix, acidic motif (VCA) segment. Multiple SH3 domain-containing adapter proteins can bind and possibly activate N-WASP, but it remains unclear how such binding events relieve autoinhibition to unmask the VCA segment and activate the Arp2/3 complex. Here, we have used purified components to reconstitute a signaling cascade driven by membrane-localized Src homology 3 (SH3) adapters and N-WASP, resulting in the assembly of dynamic actin networks. Among six SH3 adapters tested, Nck was the most potent activator of N-WASP–driven actin assembly. We identify within Nck a previously unrecognized activation motif in a linker between the first two SH3 domains. This linker sequence, reminiscent of bacterial virulence factors, directly engages the N-WASP GBD and competes with VCA binding. Our results suggest that animals, like pathogenic bacteria, have evolved peptide motifs that allosterically activate N-WASP, leading to localized actin nucleation on cellular membranes.Actin polymerization provides the force that drives membrane protrusion during cell motility as well as the propulsion of endocytic vesicles and intracellular pathogens. Branched actin networks are assembled on the surface of cellular membranes, where actin monomers are incorporated into the membrane-apposed ends of growing actin filaments (1–3). The formation of new branches is initiated by the Arp2/3 complex, which requires allosteric activation by membrane-associated nucleation-promoting factors. Neural Wiskott–Aldrich-syndrome protein (N-WASP) is an essential nucleation-promoting factor that integrates and transduces membrane-localized signals to the Arp2/3 complex.N-WASP constitutes a regulatory hub whose localization and activation state govern the spatiotemporal dynamics of actin network formation. Under resting conditions, N-WASP exists in an autoinhibited conformation in the cytoplasm. Signaling from tyrosine kinases, GTPases, and acidic phospholipids cooperatively activates N-WASP on the membrane (4–7). Two principal themes have emerged to describe N-WASP regulatory mechanisms: allosteric activation and oligomerization (8). Allosteric activation disrupts intramolecular autoinhibitory contacts between the C helix and the GTPase binding domain (GBD). These interactions maintain N-WASP in a closed conformation that sterically occludes its carboxyl-terminal verprolin-homology, connector-helix, acidic motif (VCA) segment (9, 10) (Fig. 3A). The small GTPase Cdc42 is the archetypal allosteric N-WASP activator. Cdc42 binds directly to the GBD and releases the VCA segment (11), which subsequently binds the Arp2/3 complex and promotes actin filament nucleation from the side of a preexisting actin filament. N-WASP oligomerization or clustering, mediated by signaling adapter proteins and acidic phospholipids, facilitates simultaneous interaction of two N-WASP molecules with one Arp2/3 complex. Simultaneous engagement of the constitutively inactive Arp2/3 complex by two VCA-type ligands is required for Arp2/3-mediated actin nucleation (12, 13).Open in a separate windowFig. 3.An inter-SH3 linker in Nck promotes actin network assembly by N-WASP. (A) Experimental strategy for localization of Nck to membranes containing pY₃-nephrin. (B) Nck constructs used in the deletion analysis. (C) Representative images of actin tails formed by Nck deletion constructs. Motility reactions contained 200 nM N-WASP and 100 nM Nck. (Scale bar: 5 μm.) (D) Integrated fluorescence intensity of Alexa488 actin tails. (E) Actin tail length. (F) Integrated fluorescence intensity of bead-localized Alexa568 Nck constructs. (Mean ± SD; n > 20; ***P < 0.001.) fl., Full length; PRR, proline-rich region.Src homology 3 (SH3) domain-containing adapter proteins have also been shown to activate N-WASP. Such signaling adapters often harbor multiple SH3 domains, each capable of binding a canonical polyproline motif (14). Genetic, cell biological, and biochemical evidence supports a role for the SH2/SH3 adapter protein Nck in the activation of N-WASP (15–22). Phosphorylated tyrosine residues on membrane receptors, such as the podocyte adhesion receptor nephrin and the vaccinia virus membrane protein A36R, localize Nck to the plasma membrane through its SH2 domain. Nck then directly binds N-WASP and the N-WASP–associated protein WIP, leading to localized actin polymerization (15, 20, 21, 23). Similar to Nck, Grb2 is an SH2/SH3 adapter that binds and activates N-WASP (24, 25), acting in concert with Nck to promote actin-dependent vaccinia virus motility (26). Other SH3 adapter proteins implicated in N-WASP activation include Crk-II (27), cortactin (28), Toca/CIP4 (29, 30), and Tks4/5 (31) (Fig. 1A). The mechanism by which SH3 adapters activate N-WASP is only partially understood. Although SH3-mediated oligomerization has been shown to increase N-WASP activity, these experiments were performed with a constitutively active N-WASP mutant lacking the GBD–VCA autoinhibitory interaction (32). Key unresolved questions are whether and how SH3 adapters counteract the autoinhibitory interactions that restrain access to N-WASP’s VCA segment.Open in a separate windowFig. 1.Membrane-associated actin networks assembled by SH3 adapters and N-WASP. (A) Domain organization of SH3 adapter proteins used in this study. (B) Experimental strategy for localization of His-tagged SH3 adapters using NTAD-doped membranes supported on silica microspheres. (C) Phase contrast and fluorescence images of lipid-coated beads (NTAD density: 1%, 2.5%, or 5%) incubated with the indicated SH3 adapter (250 nM), N-WASP (50 nM), Alexa488 actin, and actin regulatory components. After 15 min, reactions were fixed with glutaraldehyde. (Scale bar: 5 μm.) (D) Integrated fluorescence intensity of Alexa488 actin tails. (E) Actin tail length. (F) Integrated fluorescence intensity of bead-localized Alexa568 SH3 adapters (250 nM). (Mean ± SD; n > 20.) Asterisks indicate a significant difference between Nck1 and Grb2 at 1% and 5% NTAD: **P < 0.01; ***P < 0.001, respectively.Here, we systematically compare the ability of distinct SH3 adapters to assemble actin and Arp2/3 networks while localized to a membrane surface. To this end, we have biochemically reconstituted the N-WASP/actin signaling cascade from pure components using a membrane bilayer supported on silica microspheres. We find that membrane-localized actin assembly varies dramatically depending on the SH3 adapter, with only Nck showing robust actin assembly. Structure–function analysis revealed a previously unidentified N-WASP activation motif embedded within a 45-aa linker that connects the first two SH3 domains of Nck. This conserved inter-SH3 domain linker binds directly to the N-WASP GBD and in concert with the SH3 domains, potently stimulates actin network assembly on membranes. Nck thus uses both allosteric and oligomerization-based mechanisms to activate N-WASP.     

Detection of genomic variations and DNA polymorphisms and impact on analysis of meiotic recombination and genetic mapping

DNA polymorphisms are important markers in genetic analyses and are increasingly detected by using genome resequencing. However, the presence of repetitive sequences and structural variants can lead to false positives in the identification of polymorphic alleles. Here, we describe an analysis strategy that minimizes false positives in allelic detection and present analyses of recently published resequencing data from Arabidopsis meiotic products and individual humans. Our analysis enables the accurate detection of sequencing errors, small insertions and deletions (indels), and structural variants, including large reciprocal indels and copy number variants, from comparisons between the resequenced and reference genomes. We offer an alternative interpretation of the sequencing data of meiotic products, including the number and type of recombination events, to illustrate the potential for mistakes in single-nucleotide polymorphism calling. Using these examples, we propose that the detection of DNA polymorphisms using resequencing data needs to account for nonallelic homologous sequences.DNA polymorphisms are ubiquitous genetic variations among individuals and include single nucleotide polymorphisms (SNPs), insertions and deletions (indels), and other larger rearrangements (1–3) (Fig. 1 A and B). They can have phenotypic consequences and also serve as molecular markers for genetic analyses, facilitating linkage and association studies of genetic diseases, and other traits in humans (4–6), animals, plants, (7–10) and other organisms. Using DNA polymorphisms for modern genetic applications requires low-error, high-throughput analytical strategies. Here, we illustrate the use of short-read next-generation sequencing (NGS) data to detect DNA polymorphisms in the context of whole-genome analysis of meiotic products.Open in a separate windowFig. 1.(A) SNPs and small indels between two ecotype genomes. (B) Possible types of SVs. Col genotypes are marked in blue and Ler in red. Arrows indicate DNA segments involved in SVs between the two ecotypes. (C) Meiotic recombination events including a CO and a GC (NCO). Centromeres are denoted by yellow dots.There are many methods for detecting SNPs (11–14) and structural variants (SVs) (15–25), including NGS, which can capture nearly all DNA polymorphisms (26–28). This approach has been widely used to analyze markers in crop species such as rice (29), genes associated with diseases (6, 26), and meiotic recombination in yeast and plants (30, 31). However, accurate identification of DNA polymorphisms can be challenging, in part because short-read sequencing data have limited information for inferring chromosomal context.Genomes usually contain repetitive sequences that can differ in copy number between individuals (26–28, 31); therefore, resequencing analyses must account for chromosomal context to avoid mistaking highly similar paralogous sequences for polymorphisms. Here, we use recently published datasets to describe several DNA sequence features that can be mistaken as allelic (32, 33) and describe a strategy for differentiating between repetitive sequences and polymorphic alleles. We illustrate the effectiveness of these analyses by examining the reported polymorphisms from the published datasets.Meiotic recombination is initiated by DNA double-strand breaks (DSBs) catalyzed by the topoisomerase-like SPORULATION 11 (SPO11). DSBs are repaired as either crossovers (COs) between chromosomes (Fig. 1C), or noncrossovers (NCOs). Both COs and NCOs can be accompanied by gene conversion (GC) events, which are the nonreciprocal transfer of sequence information due to the repair of heteroduplex DNA during meiotic recombination. Understanding the control of frequency and distribution of CO and NCO (including GC) events has important implications for human health (including cancer and aneuploidy), crop breeding, and the potential for use in genome engineering. COs can be detected relatively easily by using polymorphic markers in the flanking sequences, but NCO products can only be detected if they are accompanied by a GC event. Because GCs associated with NCO result in allelic changes at polymorphic sites without exchange of flanking sequences, they are more difficult to detect. Recent advances in DNA sequencing have made the analysis of meiotic NCOs more feasible (30–32, 34); however, SVs present a challenge in these analyses. We recommend a set of guidelines for detection of DNA polymorphisms by using genomic resequencing short-read datasets. These measures improve the accuracy of a wide range of analyses by using genomic resequencing, including estimation of COs, NCOs, and GCs.     

High-resolution helix orientation in actin-bound myosin determined with a bifunctional spin label

Using electron paramagnetic resonance (EPR) of a bifunctional spin label (BSL) bound stereospecifically to Dictyostelium myosin II, we determined with high resolution the orientation of individual structural elements in the catalytic domain while myosin is in complex with actin. BSL was attached to a pair of engineered cysteine side chains four residues apart on known α-helical segments, within a construct of the myosin catalytic domain that lacks other reactive cysteines. EPR spectra of BSL-myosin bound to actin in oriented muscle fibers showed sharp three-line spectra, indicating a well-defined orientation relative to the actin filament axis. Spectral analysis indicated that orientation of the spin label can be determined within <2.1° accuracy, and comparison with existing structural data in the absence of nucleotide indicates that helix orientation can also be determined with <4.2° accuracy. We used this approach to examine the crucial ADP release step in myosin’s catalytic cycle and detected reversible rotations of two helices in actin-bound myosin in response to ADP binding and dissociation. One of these rotations has not been observed in myosin-only crystal structures.The myosin family of molecular motors is responsible for numerous vital functions in eukaryotes, including the contraction of striated muscle. Bundled within an intricate and highly regulated myofibril lattice, muscle myosin II converts the chemical energy released by ATP binding and hydrolysis into mechanical work, executing a series of structural transitions that generate force on actin and shorten each muscle cell (1, 2). Coupling of actin binding, nucleotide hydrolysis, and lever arm movement within myosin’s catalytic domain (CD) is essential for proper function of the contractile apparatus (3, 4).Myosin function requires actin, and thus an understanding of its mechanism requires analysis of both proteins in complex. However, no crystals of actin–myosin complexes have been reported, so the resolution of actin-bound myosin structures is currently limited to that of electron microscopy. Furthermore, X-ray crystallography and electron microscopy produce only static structures in frozen or crystalline environments, which cannot accurately render the dynamics, disorder, and structural transitions that are essential to understanding function and pathology (4, 5).In contrast, site-directed spectroscopy can be used to examine the actin–myosin complex under more physiological conditions. Both fluorescence and electron paramagnetic resonance (EPR) have been used in complement to examine the structural dynamics of myosin bound to actin (6, 7). EPR offers superior orientational resolution, due to the high sensitivity of the EPR spectrum to alignment of a spin label in the applied magnetic field. A well-placed spin label can provide direct information about orientation and dynamics in the vicinity of the labeling site, a strategy that has proven powerful in the study of myosin in oriented muscle fibers (7–9). However, conventional methods for site-directed spin labeling impose significant limits on the effective resolving power of EPR. Spin labels are typically incorporated into proteins through covalent attachment to Cys, resulting in a flexible linker that permits the label to undergo ns rotational motion independent of the peptide backbone. Such motion obscures the orientation dependence of the spectrum (10, 11).Our solution is to eliminate local probe motions by using a spin label that becomes strongly and stereospecifically immobilized with respect to the target protein on attachment. In site-directed fluorescence, this has been achieved by using probes that react with di-Cys (12–14) or tetra-Cys (15) labeling sites, but these fluorescent probes are typically at least twice the size of spin labels, and fluorescence lacks the high orientational resolution of EPR (6, 7). The spin-labeled amino acid TOAC provides stereospecific attachment to the peptide backbone, but this probe is currently only practical for peptides on the order of 50 amino acids or less (16). For larger proteins, spin labels have been synthesized with bulky substituents to reduce mobility (17), or substitution with additional reactive moieties to confer bifunctionality (18–20). The smallest and simplest of these derivatives shares its basic structure with the widely used methanethiosulfonate spin label [1-oxyl-2,2,5,5-tetramethyl-∆3-pyrroline-3-methyl methanethiosulfonate spin label (MTSSL)], with a second MTS group that allows bifunctional targeting of two Cys residues (20) (Fig. 1A). This bifunctional spin label (BSL) is rigidly immobilized when reacted with a pair of Cys residues, so that it undergoes negligible ns rotational motion relative to the protein and can thus be used reliably to measure μs protein rotational motions by saturation transfer EPR (21–23).Open in a separate windowFig. 1.(A) Chemical structure of BSL. (B) BSL bound stereospecifically to an α-helix at positions i and i+4, as in ref. 24. (C) Angles θ_NB and ϕ_NB that define the orientation of the nitroxide spin label (defined by axes x_N, y_N, z_N) relative to the applied magnetic field B, which directly determine the orientation dependence of the EPR spectrum. (D) Orienting the helically ordered muscle fiber (and thus the actin filament axis) with B permits direct measurement of the nitroxide orientation relative to actin.Crystallography has shown that BSL exhibits a rigid and stereospecific linkage when reacted with Cys four residues apart on successive turns of an α-helix, with great potential for accurate spin-spin distance measurements (Fig. 1B) (24). However, the potential advantages of BSL for enhanced orientational resolution have not been explored. We hypothesize that if BSL is used in the context of an intrinsically oriented system (e.g., myosin in the myofilament lattice), the resolution of EPR will be sufficient to detect the orientation of individual protein structural elements with unprecedented accuracy (Fig. 1 C and D).     

From the Cover: Phosphorylation and calcium antagonistically tune myosin-binding protein C’s structure and function

During each heartbeat, cardiac contractility results from calcium-activated sliding of actin thin filaments toward the centers of myosin thick filaments to shorten cellular length. Cardiac myosin-binding protein C (cMyBP-C) is a component of the thick filament that appears to tune these mechanochemical interactions by its N-terminal domains transiently interacting with actin and/or the myosin S2 domain, sensitizing thin filaments to calcium and governing maximal sliding velocity. Both functional mechanisms are potentially further tunable by phosphorylation of an intrinsically disordered, extensible region of cMyBP-C’s N terminus, the M-domain. Using atomic force spectroscopy, electron microscopy, and mutant protein expression, we demonstrate that phosphorylation reduced the M-domain’s extensibility and shifted the conformation of the N-terminal domain from an extended structure to a compact configuration. In combination with motility assay data, these structural effects of M-domain phosphorylation suggest a mechanism for diminishing the functional potency of individual cMyBP-C molecules. Interestingly, we found that calcium levels necessary to maximally activate the thin filament mitigated the structural effects of phosphorylation by increasing M-domain extensibility and shifting the phosphorylated N-terminal fragments back to the extended state, as if unphosphorylated. Functionally, the addition of calcium to the motility assays ablated the impact of phosphorylation on maximal sliding velocities, fully restoring cMyBP-C’s inhibitory capacity. We conclude that M-domain phosphorylation may have its greatest effect on tuning cMyBP-C’s calcium-sensitization of thin filaments at the low calcium levels between contractions. Importantly, calcium levels at the peak of contraction would allow cMyBP-C to remain a potent contractile modulator, regardless of cMyBP-C’s phosphorylation state.Cardiac contractility results from the calcium-dependent sliding of actin-based thin filaments toward the centers of myosin-based thick filaments to shorten the overall length of the sarcomere, the heart’s elementary contractile unit. Filament sliding is turned on and off on a beat-to-beat basis by calcium’s binding to, and release from, the troponin–tropomyosin regulatory complex on the thin filament. These mechanochemical regulatory processes are fine-tuned by proteins within the thick filament, including cardiac myosin-binding protein C (cMyBP-C). Although cMyBP-C is not essential for cardiac contractility, its importance in contractile function is evidenced by mutations in the MYBPC3 gene being a leading cause of inherited hypertrophic cardiomyopathy (1, 2). Because of the prevalence of this disease (affecting 1 in 500 people) and the potential for therapeutic intervention, much work during the last two decades has focused on defining cMyBP-C’s structure and function within the sarcomere (3).cMyBP-C has an elongated modular structure comprising 11 Ig and fibronectin type III (Fn3) domains, numbered C0–C10 from the N terminus (Fig. 1A) (4, 5). The C-terminal domain (C10) is tightly bound to the thick filament backbone, and the N-terminal domains extend radially from the thick filament (6, 7). Thus cMyBP-C’s N-terminal domains are positioned to bind to neighboring actin filaments and/or the myosin S2 domain to modulate actomyosin activity. However, consensus regarding the actual binding sites and partner(s) in vivo has not yet been reached (8). The N-terminal domains contain long polypeptide linkers between the C0–C1 and C1–C2 domains, the latter termed the “M-domain” or “motif” (Fig. 1A). The M-domain contains four highly conserved serines (S273, S282, S302, and S307; mouse sequence) in an intrinsically disordered region of its N-terminal half and a three-helix bundle (residues 317–351) in its C-terminal region (Fig. 1A) (9, 10). β-Adrenergic–stimulated phosphorylation of these serines is believed to enhance cardiac contractility (11, 12), and a high level of phosphorylation appears to be critical to normal cardiac function, whereas dephosphorylation has been associated with heart failure (13–15). Although the mechanistic role of cMyBP-C phosphorylation in vivo remains unclear, it is known to reduce the extensibility of the M-domain (10, 16), diminish the binding of cMyBP-C to actin (17, 18) and to myosin S2 (19), and to tune cMyBP-C’s ability to modulate actomyosin activity in vitro (20–24).Open in a separate windowFig. 1.Structure and function of cMyBP-C. (A) Schematic diagram of full-length cMyBP-C. Domains C1 and C2 are connected by the M-domain, containing an intrinsically disordered N-terminal region with four phosphorylatable serines, and a more structured C-terminal half. Ig-like domains are shown in blue, and Fn-like domains are shown in red. (B) Illustration of half of a native thick filament with a native thin filament landing on the tip of the thick filament and being translocated through the D- and C-zones at the different speeds indicated, as observed in the TIRFM experiments. (C) Wild-type C0C3 and C1C2 N-terminal fragments used in motility, AFM, and EM assays. Phosphomimetic counterparts to each fragment were expressed containing aspartic acid substitutions for the four serines highlighted in A.To gain insight into the function of cMyBP-C in its native structural environment, we recently developed a total internal reflectance microscopy (TIRFM)-based assay to visualize the sliding of actin-based filaments along native cardiac thick (myosin) filaments (21, 23). The actin filaments used in this assay are short 250-nm shards, which allow independent probing of thick filament areas with and without cMyBP-C (the C- and D-zones, respectively) (Fig. 1B). Using this assay, we showed that the sliding velocity of bare F-actin filaments was slowed within the C-zone of thick filaments and that the degree of slowing was reduced by phosphorylation of the four serines within the M-domain (21). Next, we examined the motion of native thin filaments containing the calcium-regulatory troponin–tropomyosin complex over native thick filaments. We found that cMyBP-C in the C-zone activated the native thin filaments at low calcium levels (Fig. 1B) and that the level of cMyBP-C–induced activation was lessened by phosphorylation (23). These findings collectively indicated that phosphorylated cMyBP-C is a less potent modulator of actomyosin interactions, as also has been demonstrated in other in vitro model systems (20, 22).Here, using in vitro motility assays in combination with atomic force microscopy (AFM) and electron microscopy (EM), we show that phosphorylation attenuates cMyBP-C’s function by multimodal structural ordering within the M-domain, which in turn affects long-range global interactions between cMyBP-C’s N-terminal domains. These phosphorylation-dependent structural changes suggest a molecular basis for cMyBP-C’s ability to modulate cardiac contractility. In addition, we identify another novel modulatory mechanism by which calcium limits these intra- and interdomain structural interactions and the ability of cMyBP-C phosphorylation to affect function. Our findings suggest that phosphorylation and calcium can antagonistically fine-tune cMyBP-C’s modulation of cardiac contractility, so that phosphorylation plays its greatest role when calcium levels within the sarcomere are low. The mechanistic phenomena described here may apply to additional proteins of the contractile machinery that are tuned by phosphorylation and possibly to other biological systems regulated through phosphorylation and changes in intracellular calcium levels.     

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.     

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.     

Endocytic proteins drive vesicle growth via instability in high membrane tension environment

Clathrin-mediated endocytosis (CME) is a key pathway for transporting cargo into cells via membrane vesicles; it plays an integral role in nutrient import, signal transduction, neurotransmission, and cellular entry of pathogens and drug-carrying nanoparticles. Because CME entails substantial local remodeling of the plasma membrane, the presence of membrane tension offers resistance to bending and hence, vesicle formation. Experiments show that in such high-tension conditions, actin dynamics is required to carry out CME successfully. In this study, we build on these pioneering experimental studies to provide fundamental mechanistic insights into the roles of two key endocytic proteins—namely, actin and BAR proteins—in driving vesicle formation in high membrane tension environment. Our study reveals an actin force-induced “snap-through instability” that triggers a rapid shape transition from a shallow invagination to a highly invaginated tubular structure. We show that the association of BAR proteins stabilizes vesicles and induces a milder instability. In addition, we present a rather counterintuitive role of BAR depolymerization in regulating the shape evolution of vesicles. We show that the dissociation of BAR proteins, supported by actin–BAR synergy, leads to considerable elongation and squeezing of vesicles. Going beyond the membrane geometry, we put forth a stress-based perspective for the onset of vesicle scission and predict the shapes and composition of detached vesicles. We present the snap-through transition and the high in-plane stress as possible explanations for the intriguing direct transformation of broad and shallow invaginations into detached vesicles in BAR mutant yeast cells.Clathrin-mediated endocytosis (CME) is one of the key metabolic pathways for transporting macromolecules into eukaryotic cells (1–7); it is characterized by a chain of remodeling events that transforms an almost flat patch of plasma membrane into a cargo-carrying closed vesicle. The journey from a patch to a shallow invagination, then to a mature vesicle, and finally to a detached vesicle is executed by an elaborate set of proteins that act in a well-orchestrated fashion. Because this shape evolution entails significant local bending of the membrane, it is highly sensitive to the resting tension in the membrane. A higher tension in a membrane makes a membrane taut and harder to bend, thus increasing the energetic cost required to form new vesicles. As a consequence, in cells experiencing high membrane tension, such as yeast cells and mammalian cells with polarized domains or those subjected to increased tension, actin dynamics has been found to be necessary to provide additional driving force to successfully complete CME (8–14). Although this fact has been established by seminal experimental studies, how actin forces actually drive vesicle formation and facilitate vesicle scission is not well understood. In addition, the role of another key membrane remodeling protein—the BAR protein—in overcoming tension has not yet been explored. In this paper, we pursue a detailed theoretical and computational analysis to unravel some previously unidentified mechanisms by which these key endocytic proteins (actin and BAR proteins) offset membrane tension, drive vesicle growth, and assist vesicle scission.We begin by posing a conundrum. In yeast cells, clathrin, actin, and BAR proteins contribute to vesicle formation in different capacities. Though the inhibition of actin polymerization completely arrests endocytosis (9, 11, 12, 14), the absence of clathrin and BAR proteins only leads to ∼50% and 25% reduction in the internalization events, respectively (14–18). Although a high scission rate is maintained in BAR mutant cells, there is a fundamental difference between the shape evolution process in these and the wild-type cells. In the wild-type cells, a shallow invagination turns into an elongated vesicle with a constricted neck before scission, which is successfully imaged in experimental studies (Fig. 1) (14, 18, 19). In contrast, such an intermediate shape is not observed in BAR mutant cells. After a shallow and broad invagination, experimental images directly show detached vesicles in the cytoplasm (Fig. 1) (18). This is rather intriguing because the existing model of membrane scission requires lipids to come in close proximity and pass through a hemifission state before scission to avoid any leak during the topological transition (20–23). How then does a shallow invagination directly transform into a detached vesicle? We will show in later sections that this conundrum is at the core of the shape-evolution mechanism in the presence of resting tension in the plasma membrane and is critical for understanding the roles of actin and BAR proteins in CME.Open in a separate windowFig. 1.The conundrum: In wild-type yeast cells, actin and BAR proteins turn a shallow invagination into a mature vesicle with a narrow tubular domain before scission (lipid membrane is shown in yellow, clathrin coat in red, actin filaments in blue, and BAR coat in green). In BAR mutant yeast cells, the intermediate vesicle with a constricted neck is not observed. A detached vesicle is directly seen after an initial broad invagination. Because a nonleaky scission requires lipids to come in close proximity and transition through a hemifission state, how broad and shallow invaginations undergo scission remains intriguing. This puzzle is at the core of this study. The figure is not drawn to scale.Several theoretical and computational studies have advanced our physical understanding of CME in both mammalian and yeast cells (24–27). Liu et al. (24) studied vesicle formation and scission in yeast cells under the action of curvature-generating proteins and actin filaments. The study highlighted a critical role of lipid phase boundary-induced line tension in budding and scission. In a follow-up work, temporal and spatial coordination of endocytic proteins was studied in an integrated model to simulate endocytosis in mammalian and yeast cells (25). The study showed a dynamic two-way coupling between the membrane geometry and the various biochemical reactions. Agrawal and Steigmann (26) used a unified theory of heterogeneous membrane to show that clathrin coat could drive vesicle formation without assistance from line tension in the absence of a resting plasma membrane tension. Agrawal et al. (27) studied the roles of epsin and clathrin in the nucleation of membrane vesicles. Although these studies have provided fundamental mechanistic insights into CME, the physical underpinnings of the remodeling mechanism in the presence of tension and the specific roles played by key proteins in countering tension remain unaddressed.In this study, we simulate membrane–protein interactions at the continuum scale to explore the consequences of finite tension. We first model the effect of actin forces in driving the growth of a shallow clathrin-coated vesicle. We find that until a critical force is reached, the vesicle undergoes smooth transition. Once the critical force is crossed, it experiences a snap-through transition that drastically elongates and squeezes the vesicle; this leads to a significant in-plane stress in the tubular region of the vesicle that far exceeds the rupture tension. We then model the effect of BAR proteins. We find that the attachment of BAR proteins also drives vesicle formation by instability, but it is much gentler compared with the actin case. To our surprise, we find that after the instability has occurred, the dissociation of BAR proteins leads to a larger elongation and growth of the vesicle. We predict vesicle shapes at different stages of CME that closely match those observed experimentally in yeast cells. To test the in-plane stress as a criterion for membrane scission, we simulate the geometries of detached vesicles. We find that the vesicles in the actin-driven case (in the absence of BAR proteins) are smaller than the vesicles in the BAR-driven case. In the latter case, the BAR proteins end up in the vesicle along with the clathrin coat as observed in ref. 18. Finally, we show that the membrane tension is the key parameter that regulates vesicle morphology.