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

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

TIPE2 protein serves as a negative regulator of phagocytosis and oxidative burst during infection

Phagocytosis and oxidative burst are two major effector arms of innate immunity. Although it is known that both are activated by Toll-like receptors (TLRs) and Rac GTPases, how their strengths are controlled in quiescent and TLR-activated cells is not clear. We report here that TIPE2 (TNFAIP8L2) serves as a negative regulator of innate immunity by linking TLRs to Rac. TLRs control the expression levels of TIPE2, which in turn dictates the strengths of phagocytosis and oxidative burst by binding to and blocking Rac GTPases. Consequently, TIPE2 knockout cells have enhanced phagocytic and bactericidal activities and TIPE2 knockout mice are resistant to bacterial infection. Thus, TIPE2 sets the strengths of phagocytosis and oxidative burst and may be targeted to effectively control infections.Phagocytosis and oxidative burst (or respiratory burst) are two fundamental effector mechanisms of innate immunity that work in concert to eliminate infectious microbes (1, 2). Phagocytosis allows the phagocytes of the immune system (monocytes and granulocytes) to engulf infectious microbes and to contain them in a special vacuole called a phagosome. Oxidative burst in turn injects into the vacuole reactive oxygen species (ROS) (e.g., superoxide radical and hydrogen peroxide) that kill the microbes. Deficiency in either of these innate immune mechanisms leads to immune deficiency and uncontrolled infections (3–6).Both phagocytosis and oxidative burst are controlled by the Rac proteins of the Ras small GTPase superfamily (1–4). There are three mammalian Rac GTPases, which are designated as Rac1, Rac2, and Rac3. Small GTPases are enzymes that hydrolyze GTP. They are active when bound to GTP and inactive when bound to GDP and serve as molecular “on-and-off” switches of signaling pathways that control a wide variety of cellular processes including growth, motility, vesicle trafficking, and death (7). Rac GTPases control phagocytosis by promoting actin polymerization through their effector proteins such as p21-activated kinases (PAKs), WASP family Verprolin homology domain-containing protein (WAVE), and IQ motif containing GTPase-activating protein-1 (IQGAP1) (1). Rac GTPases also mediate ROS production by binding and activating the NADPH oxidase complex through the p67(Phox) protein (1). Rac GTPase deficiency in mice and humans leads to an immune-deficient syndrome, which is characterized by defective phagocytosis and oxidative burst, recurrent infection, and granulomas (3–6).Although quiescent phagocytes are capable of phagocytosis and ROS production, their levels are low. Toll-like receptor (TLR) activation or microbial infection significantly up-regulates these innate immune processes (8–11). However, the mechanisms whereby microbes promote them are not well understood. TIPE2, or tumor necrosis factor-α–induced protein 8 (TNFAIP8)-like 2 (TNFAIP8L2), is a member of the TNFAIP8 family, which is preferentially expressed in hematopoietic cells (12–18). It is significantly down-regulated in patients with infectious or autoimmune disorders (15, 19). The mammalian TNFAIP8 family consists of four members: TNFAIP8, TIPE1, TIPE2, and TIPE3, whose functions are largely unknown (14, 20). We recently generated TIPE2-deficient mice and discovered that TIPE2 plays a crucial role in immune homeostasis (14). We report here that TIPE2 controls innate immunity by targeting the Rac GTPases.     

EFA6 controls Arf1 and Arf6 activation through a negative feedback loop

Guanine nucleotide exchange factors (GEFs) of the exchange factor for Arf6 (EFA6), brefeldin A-resistant Arf guanine nucleotide exchange factor (BRAG), and cytohesin subfamilies activate small GTPases of the Arf family in endocytic events. These ArfGEFs carry a pleckstrin homology (PH) domain in tandem with their catalytic Sec7 domain, which is autoinhibitory and supports a positive feedback loop in cytohesins but not in BRAGs, and has an as-yet unknown role in EFA6 regulation. In this study, we analyzed how EFA6A is regulated by its PH and C terminus (Ct) domains by reconstituting its GDP/GTP exchange activity on membranes. We found that EFA6 has a previously unappreciated high efficiency toward Arf1 on membranes and that, similar to BRAGs, its PH domain is not autoinhibitory and strongly potentiates nucleotide exchange on anionic liposomes. However, in striking contrast to both cytohesins and BRAGs, EFA6 is regulated by a negative feedback loop, which is mediated by an allosteric interaction of Arf6-GTP with the PH-Ct domain of EFA6 and monitors the activation of Arf1 and Arf6 differentially. These observations reveal that EFA6, BRAG, and cytohesins have unanticipated commonalities associated with divergent regulatory regimes. An important implication is that EFA6 and cytohesins may combine in a mixed negative-positive feedback loop. By allowing EFA6 to sustain a pool of dormant Arf6-GTP, such a circuit would fulfill the absolute requirement of cytohesins for activation by Arf-GTP before amplification of their GEF activity by their positive feedback loop.Guanine nucleotide exchange factors (GEFs), which activate small GTPases by stimulating their intrinsically very slow GDP/GTP exchange, are key players in the extraordinary diversity of small GTPases pathways (reviewed in ref. 1). Small GTPases carry little specificity determinants on their own to determine when and where they should be turned on and which pathway they should activate (2), which are instead largely monitored by their GEFs. Thus, understanding how different members of a GEF family activate an individual small GTPase in distinct patterns is a major issue in small GTPase biology in normal cells and in diseases.An important contribution to the functional specificity of GEFs is how they themselves are regulated. Crystallographic studies combined with biochemical studies that reconstituted GEF-stimulated GDP/GTP nucleotide exchange have been instrumental in uncovering a growing complexity of regulatory mechanisms (reviewed in ref. 1). These include autoinhibitory elements outside the catalytic GEF domain that block access to the active site (3–7), large conformational changes that release autoinhibition in response to various stimuli (8–11), positive feedback loops in which freshly produced GTP-bound GTPases stimulate GDP/GTP exchange (10, 12–15), and potentiation of nucleotide exchange by colocalization on membranes (11, 13, 16, 17).These previous studies demonstrated that a wide range of regulatory regimes can be achieved even at the scale of a single GEF family by regulatory mechanisms that combine in multiple ways. GEFs that activate small GTPases of the Arf family (ArfGEFs), which are major regulators of many aspects of membrane traffic and organelle structure in eukaryotic cells (reviewed in refs. 18 and 19), form one of the best-characterized GEF families to date (reviewed in ref. 1), making a comprehensive view of their regulatory repertoire within reach. ArfGEFs comprise two major groups: the BIG/GBF1 group, which functions at the Golgi, and a group composed of the exchange factor for Arf6 (EFA6), brefeldin A-resistant Arf guanine nucleotide exchange factor (BRAG), and cytohesin subfamilies, which activate Arf GTPases at the cell periphery and function in various aspects of endocytosis (reviewed in ref. 20). The actual substrates of these ArfGEFs have been difficult to establish, notably because the most abundant Arf isoform, Arf1, was long believed to be excluded from the plasma membrane where the Arf6 isoform is located. Accordingly, cytohesins and BRAGs have been described as Arf6-specific GEFs in cells but are now recognized as active Arf1-GEFs (16, 21, 22), whereas EFA6 remains the sole ArfGEF considered to be strictly Arf6-specific (23, 24).Members of the EFA6, BRAG, and cytohesin subfamilies have divergent N-terminal domains but a related domain organization in their C terminus comprising a Sec7 domain, which stimulates GDP/GTP exchange, followed by a pleckstrin homology (PH) domain, which has multiple regulatory functions. In cytohesins, the PH domain recognizes signaling phosphoinositides by its canonical lipid-binding site (25), autoinhibits the Sec7 domain by obstructing its Arf-binding site (4), and amplifies nucleotide exchange by a positive feedback loop involving its direct interaction with Arf1-GTP or Arf6-GTP (10, 13, 21). In contrast, the PH domain of BRAG is not autoinhibitory and is not involved in a feedback loop, but instead strongly potentiates nucleotide exchange by binding to polyanionic membranes without marked phosphoinositides preference (16).How members of the EFA6 subfamily are regulated is currently unknown. These ArfGEFs are found predominantly (although not exclusively) in the brain and function in the coordination of endocytosis and actin dynamics (23, 26, 27), in the maintenance of tight junctions (28), in microtubule dynamics in Caenorhabditis elegans embryos (29), and in the formation and maintenance of dendrites (30), although the molecular details of these functions remain largely unknown. Consistent with an important role in the brain, defects in EFA6 functions have been found in neurologic disorders (31) and in human gliomas (32). The PH domain of EFA6 subfamily members drives the localization of EFA6 members to the plasma membrane (26) and it binds to PIP₂ lipids (33). It is followed by a 150-residue C-terminal (Ct) domain predicted to form a coiled coil, which massively induces actin-rich membrane protrusions when expressed with the PH domain (26). The divergence of regulatory mechanisms between Sec7-PH–containing cytohesins and BRAGs prompted us to undertake a quantitative biochemical investigation of EFA6 nucleotide exchange regulation. Our findings reveal an overlooked dual specificity of EFA6 for Arf1 and Arf6 and an unprecedented regulation by a negative feedback loop, with important potential implications for the activation of Arf GTPases in endocytic events.     

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

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

Chronophin coordinates cell leading edge dynamics by controlling active cofilin levels

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

GluN3A expression restricts spine maturation via inhibition of GIT1/Rac1 signaling

NMDA-type glutamate receptors (NMDARs) guide the activity-dependent remodeling of excitatory synapses and associated dendritic spines during critical periods of postnatal brain development. Whereas mature NMDARs composed of GluN1 and GluN2 subunits mediate synapse plasticity and promote spine growth and stabilization, juvenile NMDARs containing GluN3A subunits are thought to inhibit these processes via yet unknown mechanisms. Here, we report that GluN3A binds G protein-coupled receptor kinase-interacting protein (GIT1), a postsynaptic scaffold that assembles actin regulatory complexes, including the Rac1 guanine nucleotide exchange factor βPIX, to promote Rac1 activation in spines. Binding to GluN3A limits the synaptic localization of GIT1 and its ability to complex βPIX, leading to decreased Rac1 activation and reduced spine density and size in primary cultured neurons. Conversely, knocking out GluN3A favors the formation of GIT1/βPIX complexes and increases the activation of Rac1 and its main effector p21-activated kinase. We further show that binding of GluN3A to GIT1 is regulated by synaptic activity, a response that might restrict the negative regulatory effects of GluN3A on actin signaling to inactive synapses. Our results identify inhibition of Rac1/p21-activated kinase actin signaling pathways as an activity-dependent mechanism mediating the inhibitory effects of GluN3A on spine morphogenesis.During the development of neural circuits, a phase of intense synaptogenesis is followed by a period of activity-dependent remodeling (or “synaptic refinement”) in which more than half of the initially formed synapses are eliminated, whereas other connections will mature and be kept (1, 2). The subunit composition of NMDA-type glutamate receptors (NMDARs) expressed by individual synapses during this critical period is a key factor influencing functional and structural synaptic plasticity and, in turn, synapse fate (3). Mature NMDARs composed of GluN1 and GluN2 subunits drive the maturation of active synapses by detecting coincident pre- and postsynaptic activity and coupling this activity to signaling pathways that trigger the enlargement and stabilization of synapses and associated dendritic spines (4–6). This structural plasticity is critical for coupling the wiring of neural circuits to experience and supporting the long-term maintenance of spines and memories. During the refinement stage, NMDARs additionally contain GluN3A subunits that serve as a brake on synapse maturation and stabilization, which might provide a counterbalance to limit synapse numbers. Supporting this idea, loss of GluN3A increases spine density and size (7) and accelerates the expression of markers of synaptic maturation (8), whereas overexpression reduces synapse and spine density and yields a higher proportion of smaller, immature spines (9). However, the downstream mechanisms by which GluN3A inhibits synapse and spine maturation remain unknown.Spines are actin-rich, and their structural remodeling relies on rearrangements of the actin cytoskeleton (10–13). Cytoskeletal rearrangements are regulated by the Rho family of small GTPases, and two members of this family, Rac1 and RhoA, are major regulators of spine remodeling (14, 15). Rho-GTPases act as molecular switches that cycle between an inactive GDP-bound conformation and an active GTP-bound conformation (16). Their activation state is controlled by guanine exchange factors (GEFs), which promote the exchange of GDP for GTP, and GTPase-activating proteins (GAPs), which catalyze GTP hydrolysis. Several Rac1-specific GEFs, including Kalirin7, Tiam1, and βPIX, are targeted to synapses via interactions with scaffolding proteins, which allows local regulation of actin remodeling in spines and its coupling to synaptic activity (12, 17–22). Although many studies have shown that NMDAR activation induces cytoskeletal and spine remodeling by activating Rac1-GEFs (23–25), much less is known about pathways that restrict excitatory synapse maturation and/or promote elimination.Here, we identify a physical association between the intracellular C-terminal domain of GluN3A subunits and G protein-coupled receptor kinase-interacting protein (GIT1), a postsynaptic scaffold that assembles a multiprotein signaling complex with Rac1 and the Rac1-GEF βPIX to regulate actin dynamics in spines (19, 26). GIT1 selectively bound juvenile NMDARs containing GluN3A but not mature NMDAR subtypes, and binding was regulated by activity because it could be enhanced or reduced, respectively, by brief episodes of synaptic inactivity or synaptic stimulation. A functional analysis demonstrates that binding to GluN3A interferes with the synaptic localization of GIT1 and its ability to recruit βPIX, leading to decreased Rac1 activation. We finally show that GluN3A-induced reductions in spine density and size critically require GIT1 binding. We propose that the coupling of GIT1/GluN3A binding to synapse use might provide an effective mechanism with which to restrict the maturation and growth of inactive synapses in a selective manner.     

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.     

Arf guanine nucleotide-exchange factors BIG1 and BIG2 regulate nonmuscle myosin IIA activity by anchoring myosin phosphatase complex

Brefeldin A-inhibited guanine nucleotide-exchange factors BIG1 and BIG2 activate, through their Sec7 domains, ADP ribosylation factors (Arfs) by accelerating the replacement of Arf-bound GDP with GTP for initiation of vesicular transport or activation of specific enzymes that modify important phospholipids. They are also implicated in regulation of cell polarization and actin dynamics for directed migration. Reciprocal coimmunoprecipitation of endogenous HeLa cell BIG1 and BIG2 with myosin IIA was demonstrably independent of Arf guanine nucleotide-exchange factor activity, because effects of BIG1 and BIG2 depletion were reversed by overexpression of the cognate BIG molecule C-terminal sequence that follows the Arf activation site. Selective depletion of BIG1 or BIG2 enhanced specific phosphorylation of myosin regulatory light chain (T18/S19) and F-actin content, which impaired cell migration in Transwell assays. Our data are clear evidence of these newly recognized functions for BIG1 and BIG2 in transduction or integration of mechanical signals from integrin adhesions and myosin IIA-dependent actin dynamics. Thus, by anchoring or scaffolding the assembly, organization, and efficient operation of multimolecular myosin phosphatase complexes that include myosin IIA, protein phosphatase 1δ, and myosin phosphatase-targeting subunit 1, BIG1 and BIG2 serve to integrate diverse biophysical and biochemical events in cells.Cell migration requires the coordinated spatiotemporal regulation of actomyosin function for alterations in cell shape and adhesion. Nonmuscle myosin II (NM II) is critical for regulation of structural remodeling and migration of nonmuscle cells. NM II comprises two heavy chains of 230 kDa, two 20-kDa regulatory light chains (RLCs), and two 17-kDa essential light chains that assemble into bipolar filaments with actin-stimulated ATPase activity. The resultant contractility and actin cross-linking drive assembly of actin filaments that form the actin cytoskeleton (1). In mammalian cells, NM II heavy chain proteins (NMHC) IIA, IIB, and IIC encoded, respectively, by three genes (Myh9, Myh10, and Myh14), are 60–80% identical. Three hexameric isoforms of NM II named NM IIA, IIB, and IIC, with both shared and unique properties, are designated by NMHC component, which accounts for their differences, and all function in events like cell polarization, migration, and adhesion that involve mechano-sensing and motility (2, 3).Reversible phosphorylation of specific amino acids in the pair of RLCs and/or the heavy chains alters NM II activity. Without phosphorylation, NM II folds into a compact structure, in which one head interacts with the second head of the same molecule. The tails interact also with the heads to prevent ATP hydrolysis and thereby filament assembly. Phosphorylation of RLC on T18 and/or S19 disrupts head–head and head–tail interactions and promotes the formation of contractile actin bundles by enhancing the actin-activated ATPase of NM II (1). Rho-associated protein kinase (ROCK) (4), myotonic dystrophy kinase-related Cdc42-binding kinase (5), and myosin light chain kinase (MLCK) (6), the kinases most widely implicated in RLC T18 and/or S19 phosphorylation, are controlled by different signaling pathways. MLCK was activated by Ca2⁺-calmodulin (6), whereas the small GTPase RhoA activated ROCK (4). Dephosphorylation of RLC T18 and/or S19 was catalyzed by myosin phosphatase, a heterotrimeric enzyme comprising the 130-kDa myosin phosphatase-targeting subunit 1 (MYPT1), the 37-kDa catalytic subunit of protein phosphatase type 1δ (PP1cδ), and a 20-kDa subunit of unknown function (7). Myosin phosphatase and NM II are associated by direct interaction of the latter with both the PP1cδ and MYPT (8–11). Phosphorylation of MYPT was reported to alter the interaction of myosin phosphatase with NM II (7, 12) and myosin phosphatase catalytic activity (7, 13, 14).Brefeldin A-inhibited guanine nucleotide-exchange proteins (BIG)1 (∼200 kDa) and BIG2 (∼190 kDa), which were initially purified together in a ∼670-kDa complex from bovine brain cytosol (15–17), like other members of the Sec7 family of ADP ribosylation factor (Arf) guanine nucleotide-exchange factors (GEFs), accelerate replacement of Arf-bound GDP with GTP to generate active Arf-GTP. Association of Arf-GTP with membranes, along with coat proteins and adaptors from cytosol, initiates vesicle formation for transport of cargo between Golgi and plasma membrane (PM) (15, 17–19). Microscopically, BIG1 was seen concentrated at trans-Golgi network, where it partially overlapped BIG2, which was associated also with recycling endosomes (19–21). Functions of BIG1 and BIG2 at the trans-Golgi network were described as redundant (22), although each protein clearly has specific roles that are not shared with the other. BIG1 was required for Golgi integrity, correct glycosylation of mature integrin β1 (23), and directed migration during wound healing (24). In contrast, BIG2 was involved in recycling of internalized transferrin receptors (21) and integrin β1 (25) to the cell surface, and in regulation of tumor necrosis factor receptor release in exosome-like vesicles (26). Besides the catalytic GEF activity of the Sec7 domain, there are additional highly conserved domains and sequences in BIG1 and BIG2 that remain functionally less well defined (27). They may act as scaffolds or platforms for assembly of multimolecular complexes required for integration of diverse signals that regulate several cellular processes. Both BIG1 and BIG2 molecules contain A kinase-anchoring protein sequences in their N-terminal region that bind specific regulatory subunits of protein kinase A (PKA) and act as scaffolds for assembly of multimolecular machines that limit intracellular PKA-cAMP signaling in time and space (28). PP1γ (29) and phosphodiesterase 3A (30) that, respectively, dephosphorylate PKA-modified BIG1 and BIG2 and terminate cAMP signals, have been identified in complexes with BIG1 and/or BIG2.Depletion of BIG1 or BIG2 was reported to decrease cell motility as well as actin-based membrane protrusions (24, 25), although the mechanisms through which these proteins altered cytoskeleton dynamics remain unclear. Here, we provide evidence that NM IIA activity in HeLa cells, which is crucial for actin stress fiber formation, is also influenced by previously unrecognized interactions with the C termini of BIG1 and BIG2, independent of Arf GEF activity, in complexes that include PP1cδ, and MYPT1.     

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

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

Dissecting neural pathways for forgetting in Drosophila olfactory aversive memory

Recent studies have identified molecular pathways driving forgetting and supported the notion that forgetting is a biologically active process. The circuit mechanisms of forgetting, however, remain largely unknown. Here we report two sets of Drosophila neurons that account for the rapid forgetting of early olfactory aversive memory. We show that inactivating these neurons inhibits memory decay without altering learning, whereas activating them promotes forgetting. These neurons, including a cluster of dopaminergic neurons (PAM-β′1) and a pair of glutamatergic neurons (MBON-γ4>γ1γ2), terminate in distinct subdomains in the mushroom body and represent parallel neural pathways for regulating forgetting. Interestingly, although activity of these neurons is required for memory decay over time, they are not required for acute forgetting during reversal learning. Our results thus not only establish the presence of multiple neural pathways for forgetting in Drosophila but also suggest the existence of diverse circuit mechanisms of forgetting in different contexts.Although forgetting commonly has a negative connotation, it is a functional process that shapes memory and cognition (1–4). Recent studies, including work in relatively simple invertebrate models, have started to reveal basic biological mechanisms underlying forgetting (5–15). In Drosophila, single-session Pavlovian conditioning by pairing an odor (conditioned stimulus, CS) with electric shock (unconditioned stimulus, US) induces aversive memories that are short-lasting (16). The memory performance of fruit flies is observed to drop to a negligible level within 24 h, decaying rapidly early after training and slowing down thereafter (17). Memory decay or forgetting requires the activation of the small G protein Rac, a signaling protein involved in actin remodeling, in the mushroom body (MB) intrinsic neurons (6). These so-called Kenyon cells (KCs) are the neurons that integrate CS–US information (18, 19) and support aversive memory formation and retrieval (20–22). In addition to Rac, forgetting also requires the DAMB dopamine receptor (7), which has highly enriched expression in the MB (23). Evidence suggests that the dopamine-mediated forgetting signal is conveyed to the MB by dopamine neurons (DANs) in the protocerebral posterior lateral 1 (PPL1) cluster (7, 24). Therefore, forgetting of olfactory aversive memory in Drosophila depends on a particular set of intracellular molecular pathways within KCs, involving Rac, DAMB, and possibly others (25), and also receives modulation from extrinsic neurons. Although important cellular evidence supporting the hypothesis that memory traces are erased under these circumstances is still lacking, these findings lend support to the notion that forgetting is an active, biologically regulated process (17, 26).Although existing studies point to the MB circuit as essential for forgetting, several questions remain to be answered. First, whereas the molecular pathways for learning and forgetting of olfactory aversive memory are distinct and separable (6, 7), the neural circuits seem to overlap. Rac-mediated forgetting has been localized to a large population of KCs (6), including the γ-subset, which is also critical for initial memory formation (21, 27). The site of action of DAMB for forgetting has yet to be established; however, the subgroups of PPL1-DANs implicated in forgetting are the same as those that signal aversive reinforcement and are required for learning (28–30). It leaves open the question of whether the brain circuitry underlying forgetting and learning is dissociable, or whether forgetting and learning share the same circuit but are driven by distinct activity patterns and molecular machinery (26). Second, shock reinforcement elicits multiple memory traces through at least three dopamine pathways to different subdomains in the MB lobes (28, 29). Functional imaging studies have also revealed Ca²⁺-based memory traces in different KC populations (31). It is poorly understood how forgetting of these memory traces differs, and it remains unknown whether there are multiple regulatory neural pathways. Notably, when PPL1-DANs are inactivated, forgetting still occurs, albeit at a lower rate (7). This incomplete block suggests the existence of an additional pathway(s) that conveys forgetting signals to the MB. Third, other than memory decay over time, forgetting is also observed through interference (32, 33), when new learning or reversal learning is introduced after training (6, 34, 35). Time-based and interference-based forgetting shares a similar dependence on Rac and DAMB (6, 7). However, it is not known whether distinct circuits underlie forgetting in these different contexts.In the current study, we focus on the diverse set of MB extrinsic neurons (MBENs) that interconnect the MB lobes with other brain regions, which include 34 MB output neurons (MBONs) of 21 types and ∼130 dopaminergic neurons of 20 types in the PPL1 and protocerebral anterior medial (PAM) clusters (36, 37). These neurons have been intensively studied in olfactory memory formation, consolidation, and retrieval in recent years (e.g., 24, 28–30, 38–48); however, their roles in forgetting have not been characterized except for the aforementioned PPL1-DANs. In a functional screen, we unexpectedly found that several Gal4 driver lines of MBENs showed significantly better 3-h memory retention when the Gal4-expressing cells were inactivated. The screen has thus led us to identify two types of MBENs that are not involved in initial learning but play important and additive roles in mediating memory decay. Furthermore, neither of these MBEN types is required for reversal learning, supporting the notion that there is a diversity of neural circuits that drive different forms of forgetting.     

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.     

Helical buckling of actin inside filopodia generates traction

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

From the Cover: PNAS Plus: Arl1p regulates spatial membrane organization at the trans-Golgi network through interaction with Arf-GEF Gea2p and flippase Drs2p

ADP ribosylation factors (Arfs) are the central regulators of vesicle trafficking from the Golgi complex. Activated Arfs facilitate vesicle formation through stimulating coat assembly, activating lipid-modifying enzymes and recruiting tethers and other effectors. Lipid translocases (flippases) have been implicated in vesicle formation through the generation of membrane curvature. Although there is no evidence that Arfs directly regulate flippase activity, an Arf-guanine-nucleotide-exchange factor (GEF) Gea2p has been shown to bind to and stimulate the activity of the flippase Drs2p. Here, we provide evidence for the interaction and activation of Drs2p by Arf-like protein Arl1p in yeast. We observed that Arl1p, Drs2p and Gea2p form a complex through direct interaction with each other, and each interaction is necessary for the stability of the complex and is indispensable for flippase activity. Furthermore, we show that this Arl1p-Drs2p-Gea2p complex is specifically required for recruiting golgin Imh1p to the Golgi. Our results demonstrate that activated Arl1p can promote the spatial modulation of membrane organization at the trans-Golgi network through interacting with the effectors Gea2p and Drs2p.ADP ribosylation factors (Arf)/SARs, a subfamily of the Ras small GTP-binding protein superfamily, are critical regulators of vesicular trafficking in eukaryotic cells (1–3). Like Ras, Arf and Arf-like (Arl) proteins cycle between the inactive GDP-bound form and active GTP-bound forms to carry out their functions. The conversion from the GDP-bound to the GTP-bound form is facilitated by Arf guanine-nucleotide-exchange factor (Arf-GEF), whereas GTP hydrolysis is catalyzed by an Arf GTPase-activating protein (Arf-GAP) (4, 5). Activated Arf recruits numerous proteins to the membrane or activates lipid-modifying enzymes to facilitate vesicle formation, including vesicle coat assembly, membrane curvature generation, lipid composition alteration, and final membrane fission (4, 6–8).Arl1p is the best-studied Arl protein and is expressed in organisms ranging from yeast to humans (9, 10). Both yeast Arl1p and human Arl1 localize to the trans-Golgi network (TGN), where they regulate multiple membrane-trafficking pathways (9, 11, 12). Yeast Arl1p participates in three different pathways at the TGN, including glycosylphosphatidylinositol (GPI)-anchored protein transport, clathrin adaptor protein Gga recruitment, and targeting of the GRIP domain–containing protein Imh1p to the Golgi. In addition to Imh1p, the Arf-GEF–like protein, Mon2p (also called Ysl2p), and the clathrin adaptor protein, Gga2p, directly interact with Arl1p and are considered to be Arl1p effectors (13).Arf-GEFs catalyze the GTP binding of Arf/Arl through the conserved Sec7 domain and are upstream regulators of Arf/Arl activities (14, 15). In yeast, the human GBF1 homolog Gea2p, which functions redundantly with Gea1p, displays Arf-GEF activity toward Arf1p and is believed to be involved in ER-Golgi and intra-Golgi transport (16, 17). Recently, Gea2p was shown to directly interact with the TGN-localized aminophospholipid translocase Drs2p and to stimulate its flippase activity (18, 19). Similarly, it was reported that the Arf-GEF–like protein Mon2p interacted with both Arl1p and the Drs2p family protein Neo1p and that the Mon2-Neo1-Arl1 complex cooperated to regulate the Golgi recruitment of Gga2p (13). These results suggest that Arf-GEFs may act with members of the Drs2p subfamily of P-type ATPases to facilitate lipid translocation. However, the role of Gea2p in Drs2p-mediated flippase activity is not clear.Flippases translocate specific phospholipid molecules to establish the asymmetry of membranes (19–22). Therefore, flippases can drive membrane bending toward the cytosol and contribute to vesicle-mediated protein transport by facilitating the generation of membrane curvature that is inherent to this process (23). The inactivation of Drs2p results in defects in the formation of an Arf/clathrin-dependent class of post-Golgi vesicles that carry exocytic cargo (24). Moreover, a mutation in DRS2 was shown to cause synthetic lethality with clathrin heavy chain and ARF1 (25). These observations indicate the importance of flippase activity for vesicle trafficking, and Drs2p is involved in protein transport at the TGN.To explore the possibility that Arl1p might, like Arf proteins, play a role in regulating membrane dynamics, we use an analysis of both physical interaction and functional interaction to demonstrate that Drs2p and Gea2p are effectors of Arl1p. We find that these proteins—Arl1p, Drs2p, and Gea2p—are all required to form a stable complex to stimulate Drs2p flippase activity and complete certain functions of Arl1p. Our study provides insight into the molecular mechanisms of spatial membrane dynamics at the TGN and their regulation by the Arl1p-Gea2p-Drs2p cooperating network.     

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

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

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

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

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

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

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.     

Mitigation of acute kidney injury by cell-cycle inhibitors that suppress both CDK4/6 and OCT2 functions

Acute kidney injury (AKI) is a potentially fatal syndrome characterized by a rapid decline in kidney function caused by ischemic or toxic injury to renal tubular cells. The widely used chemotherapy drug cisplatin accumulates preferentially in the renal tubular cells and is a frequent cause of drug-induced AKI. During the development of AKI the quiescent tubular cells reenter the cell cycle. Strategies that block cell-cycle progression ameliorate kidney injury, possibly by averting cell division in the presence of extensive DNA damage. However, the early signaling events that lead to cell-cycle activation during AKI are not known. In the current study, using mouse models of cisplatin nephrotoxicity, we show that the G1/S-regulating cyclin-dependent kinase 4/6 (CDK4/6) pathway is activated in parallel with renal cell-cycle entry but before the development of AKI. Targeted inhibition of CDK4/6 pathway by small-molecule inhibitors palbociclib (PD-0332991) and ribociclib (LEE011) resulted in inhibition of cell-cycle progression, amelioration of kidney injury, and improved overall survival. Of additional significance, these compounds were found to be potent inhibitors of organic cation transporter 2 (OCT2), which contributes to the cellular accumulation of cisplatin and subsequent kidney injury. The unique cell-cycle and OCT2-targeting activities of palbociclib and LEE011, combined with their potential for clinical translation, support their further exploration as therapeutic candidates for prevention of AKI.Cell division is a fundamental biological process that is tightly regulated by evolutionarily conserved signaling pathways (1, 2). The initial decision to start cell division, the fidelity of subsequent DNA replication, and the final formation of daughter cells is monitored and regulated by these essential pathways (2–6). The cyclin-dependent kinases (CDKs) are the central players that orchestrate this orderly progression through the cell cycle (1, 2, 6, 7). The enzymatic activity of CDKs is regulated by complex mechanisms that include posttranslational modifications and expression of activating and inhibitory proteins (1, 2, 6, 7). The spatial and temporal changes in the activity of these CDK complexes are thought to generate the distinct substrate specificities that lead to sequential and unidirectional progression of the cell cycle (1, 8, 9).Cell-cycle deregulation is a universal feature of human cancer and a long-sought-after target for anticancer therapy (1, 10–13). Frequent genetic or epigenetic changes in mitogenic pathways, CDKs, cyclins, or CDK inhibitors are observed in various human cancers (1, 4, 11). In particular, the G1/S-regulating CDK4/6–cyclin D–inhibitors of CDK4 (INK4)–retinoblastoma (Rb) protein pathway frequently is disrupted in cancer cells (11, 14). These observations provided an impetus to develop CDK inhibitors as anticancer drugs. However, the earlier class of CDK inhibitors had limited specificity, inadequate clinical activity, poor pharmacokinetic properties, and unacceptable toxicity profiles (10, 11, 14, 15). These disappointing initial efforts now have been followed by the development of the specific CDK4/6 inhibitors palbociclib (PD0332991), ribociclib (LEE011), and abemaciclib (LY2835219), which have demonstrated manageable toxicities, improved pharmacokinetic properties, and impressive antitumor activity, especially in certain forms of breast cancer (14, 16). Successful early clinical trials with these three CDK4/6 inhibitors have generated cautious enthusiasm that these drugs may emerge as a new class of anticancer agents (14, 17). Palbociclib recently was approved by Food and Drug Administration for the treatment of metastatic breast cancer and became the first CDK4/6 inhibitor approved for anticancer therapy (18).In addition to its potential as an anticancer strategy, CDK4/6 inhibition in normal tissues could be exploited therapeutically for wide-ranging clinical conditions. For example, radiation-induced myelosuppression, caused by cell death of proliferating hematopoietic stem/progenitor cells, can be rescued by palbociclib (19, 20). Furthermore, cytotoxic anticancer agents cause significant toxicities to normal proliferating cells, which possibly could be mitigated by the concomitant use of CDK4/6 inhibitors (20, 21). More broadly, cell-cycle inhibition could have beneficial effects in disorders in which maladaptive proliferation of normal cells contributes to the disease pathology, as observed in vascular proliferative diseases, hyperproliferative skin diseases, and autoimmune disorders (22, 23). In support of this possibility, palbociclib treatment recently was reported to ameliorate disease progression in animal models of rheumatoid arthritis through cell-cycle inhibition of synovial fibroblasts (24).Abnormal cellular proliferation also is a hallmark of various kidney diseases (25), and cell-cycle inhibition has been shown to ameliorate significantly the pathogenesis of polycystic kidney disease (26), nephritis (27), and acute kidney injury (AKI) (28). Remarkably, during AKI, the normally quiescent renal tubular cells reenter the cell cycle (29–34), and blocking cell-cycle progression can reduce renal injury (28). Here, we provide evidence that the CDK4/6 pathway is activated early during AKI and demonstrate significant protective effects of CDK4/6 inhibitors in animal models of cisplatin-induced AKI. In addition, we found that the CDK4/6 inhibitors palbociclib and LEE011 are potent inhibitors of organic cation transporter 2 (OCT2), a cisplatin uptake transporter highly expressed in renal tubular cells (35–37). Our findings provide a rationale for the clinical development of palbociclib and LEE011 for the prevention and treatment of AKI.