Differential Regulation of Actin Polymerization and Structure by Yeast Formin Isoforms

The budding yeast formins, Bnr1 and Bni1, behave very differently with respect to their interactions with muscle actin. However, the mechanisms underlying these differences are unclear, and these formins do not interact with muscle actin in vivo. We use yeast wild type and mutant actins to further assess these differences between Bnr1 and Bni1. Low ionic strength G-buffer does not promote actin polymerization. However, Bnr1, but not Bni1, causes the polymerization of pyrene-labeled Mg-G-actin in G-buffer into single filaments based on fluorometric and EM observations. Polymerization by Bnr1 does not occur with Ca-G-actin. By cosedimentation, maximum filament formation occurs at a Bnr1:actin ratio of 1:2. The interaction of Bnr1 with pyrene-labeled S265C Mg-actin yields a pyrene excimer peak, from the cross-strand interaction of pyrene probes, which only occurs in the context of F-actin. In F-buffer, Bnr1 promotes much faster yeast actin polymerization than Bni1. It also bundles the F-actin in contrast to the low ionic strength situation where only single filaments form. Thus, the differences previously observed with muscle actin are not actin isoform-specific. The binding of both formins to F-actin saturate at an equimolar ratio, but only about 30% of each formin cosediments with F-actin. Finally, addition of Bnr1 but not Bni1 to pyrene-labeled wild type and S265C Mg-F actins enhanced the pyrene- and pyrene-excimer fluorescence, respectively, suggesting Bnr1 also alters F-actin structure. These differences may facilitate the ability of Bnr1 to form the actin cables needed for polarized delivery of nutrients and organelles to the growing yeast bud.Bni1 and Bnr1 are the two formin isoforms expressed in Saccharomyces cerevisiae (1, 2). These proteins, as other isoforms in the formin family, are large multidomain proteins (3, 4). Several regulatory domains, including one for binding the G-protein rho, are located at the N-terminal half of the protein (4–7). FH1, FH2, and Bud6 binding domains are located in the C-terminal half of the protein (8). The formin homology 1 (FH1)² domain contains several sequential poly-l-proline motifs, and it interacts with the profilin/actin complex to recruit actin monomers and regulate the insertion of actin monomers at the barbed end of actin (9–11). The fomin homology domain 2 (FH2) forms a donut-shaped homodimer, which wraps around actin dimers at the barbed end of actin filaments (12, 13). One important function of formin is to facilitate actin polymerization by stabilizing actin dimers or trimers under polymerization conditions and then to processively associate with the barbed end of the elongating filament to control actin filament elongation kinetics (13–18).A major unsolved protein in the study of formins is the elucidation of the individual functions of different isoforms and their regulation. In vivo, these two budding yeast formins have distinct cellular locations and dynamics (1, 2, 19, 20). Bni1 concentrates at the budding site before the daughter cell buds from the mother cell, moves along with the tip of the daughter cell, and then travels back to the neck between daughter and mother cells at the end of segregation. Bnr1 localizes only at the neck of the budding cell in a very short period of time after bud emergence. Although a key cellular function of these two formins in yeast is to promote actin cable formation (8, 18), the roles of the individual formins in different cellular process is unclear because deleting either individual formin gene has limited impact on cell growth and deleting both genes together is lethal (21).Although each of the two formins can nucleate actin filament formation in vitro, the manner in which they affect polymerization is distinctly isoform-specific. Most of this mechanistic work in vitro has used formin fragments containing the FH1 and FH2 domains. Bni1 alone processively caps the barbed end of actin filaments partially inhibiting polymerization at this end (14, 16, 18). The profilin-actin complex, recruited to the actin barbed end through its binding to Bni1 FH1 domain, possibly raises the local actin concentration and appears to allow this inhibition to be overcome, thereby, accelerating barbed end polymerization. It has also been shown that this complex modifies the kinetics of actin dynamics at the barbed end (9, 11, 18, 22). Moreover, Bni1 participation leads only to the formation of single filaments (8). In comparison, the Bnr1 FH1-FH2 domain facilitates actin polymerization much more efficiently than does Bni1. Moseley and Goode (8) showed Bnr1 accelerates actin polymerization up to 10 times better than does Bni and produces actin filament bundles when the Bnr1/actin molar ratio is above 1:2. Finally, the regulation of Bni1 and Bnr1 by formin binding is different. For example, Bud 6/Aip3, a yeast cell polarity factor, binds to Bni1, but not Bnr1, and also stimulates its activity in vitro.For their studies, Moseley and Goode (8) utilized mammalian skeletal muscle actin instead of the S. cerevisiae actin with which the yeast formins are designed to function. It is entirely possible that the differences observed with the two formins are influenced quantitatively or qualitatively by the nature of the actin used in the study. This possibility must be seriously considered because although yeast and muscle actins are 87% identical in sequence, they display marked differences in their polymerization behavior (23). Yeast actin nucleates filaments better than muscle actin (24, 25). It appears to form shorter and more flexible filaments than does muscle actin (26, 27). Finally, the disposition of the P_i released during the hydrolysis of ATP that occurs during polymerization is different. Yeast actin releases its P_i concomitant with hydrolysis of the bound ATP whereas muscle actin retains the P_i for a significant amount of time following nucleotide hydrolysis (28, 29). This difference is significant because ADP-P_i F-actin has been shown to be more stable than ADP F-actin (30). Another example of this isoform dependence is the interaction of yeast Arp2/3 with yeast versus muscle actins (31). Yeast Arp2/3 complex accelerates polymerization of muscle actin only in the presence of a nucleation protein factor such as WASP. However, with yeast actin, no such auxiliary protein is required. In light of these actin behavioral differences, to better understand the functional differences of these two formins in vivo, we have studied the behavior of Bni 1 and Bnr 1 with WT and mutant yeast actins, and we have also explored the molecular basis underlying the Bnr 1-induced formation of actin nuclei from G-actin.     

F-Actin Dynamics in Neurospora crassa

This study demonstrates the utility of Lifeact for the investigation of actin dynamics in Neurospora crassa and also represents the first report of simultaneous live-cell imaging of the actin and microtubule cytoskeletons in filamentous fungi. Lifeact is a 17-amino-acid peptide derived from the nonessential Saccharomyces cerevisiae actin-binding protein Abp140p. Fused to green fluorescent protein (GFP) or red fluorescent protein (TagRFP), Lifeact allowed live-cell imaging of actin patches, cables, and rings in N. crassa without interfering with cellular functions. Actin cables and patches localized to sites of active growth during the establishment and maintenance of cell polarity in germ tubes and conidial anastomosis tubes (CATs). Recurrent phases of formation and retrograde movement of complex arrays of actin cables were observed at growing tips of germ tubes and CATs. Two populations of actin patches exhibiting slow and fast movement were distinguished, and rapid (1.2 μm/s) saltatory transport of patches along cables was observed. Actin cables accumulated and subsequently condensed into actin rings associated with septum formation. F-actin organization was markedly different in the tip regions of mature hyphae and in germ tubes. Only mature hyphae displayed a subapical collar of actin patches and a concentration of F-actin within the core of the Spitzenkörper. Coexpression of Lifeact-TagRFP and β-tubulin–GFP revealed distinct but interrelated localization patterns of F-actin and microtubules during the initiation and maintenance of tip growth.Actins are highly conserved proteins found in all eukaryotes and have an enormous variety of cellular roles. The monomeric form (globular actin, or G-actin) can self-assemble, with the aid of numerous actin-binding proteins (ABPs), into microfilaments (filamentous actin, or F-actin), which, together with microtubules, form the two major components of the fungal cytoskeleton. Numerous pharmacological and genetic studies of fungi have demonstrated crucial roles for F-actin in cell polarity, exocytosis, endocytosis, cytokinesis, and organelle movement (6, 7, 20, 34, 35, 51, 52, 59). Phalloidin staining, immunofluorescent labeling, and fluorescent-protein (FP)-based live-cell imaging have revealed three distinct subpopulations of F-actin-containing structures in fungi: patches, cables, and rings (1, 14, 28, 34, 60, 63, 64). Actin patches are associated with the plasma membrane and represent an accumulation of F-actin around endocytic vesicles (3, 26, 57). Actin cables are bundles of actin filaments stabilized with cross-linking proteins, such as tropomyosins and fimbrin, and are assembled by formins at sites of active growth, where they form tracks for myosin V-dependent polarized secretion and organelle transport (10, 16, 17, 27, 38, 47, 48). Cables, unlike patches, are absolutely required for polarized growth in the budding yeast Saccharomyces cerevisiae (34, 38). Contractile actomyosin rings are essential for cytokinesis in budding yeast, whereas in filamentous fungi, actin rings are less well studied but are known to be involved in septum formation (20, 28, 34, 39, 40).Actin cables and patches have been particularly well studied in budding yeast. However, there are likely to be important differences between F-actin architecture and dynamics in budding yeast and those in filamentous fungi, as budding yeasts display only a short period of polarized growth during bud formation, which is followed by isotropic growth over the bud surface (10). Sustained polarized growth during hyphal morphogenesis is a defining feature of filamentous fungi (21), making them attractive models for studying the roles of the actin cytoskeleton in cell polarization, tip growth, and organelle transport.In Neurospora crassa and other filamentous fungi, disruption of the actin cytoskeleton leads to rapid tip swelling, which indicates perturbation of polarized tip growth, demonstrating a critical role for F-actin in targeted secretion to particular sites on the plasma membrane (7, 22, 29, 56). Immunofluorescence studies of N. crassa have shown that F-actin localizes to hyphal tips as “clouds” and “plaques” (7, 54, 59). However, immunolabeling has failed to reveal actin cables in N. crassa and offers limited insights into F-actin dynamics. Live-cell imaging of F-actin architecture and dynamics has not been accomplished in N. crassa, yet it is expected to yield key insights into cell polarization, tip growth, and intracellular transport.We took advantage of a recently developed live-cell imaging probe for F-actin called Lifeact (43). Lifeact is a 17-amino-acid peptide derived from the N terminus of the budding yeast actin-binding protein Abp140 (5, 63) and has recently been demonstrated to be a universal live-cell imaging marker for F-actin in eukaryotes (43). Here, we report the successful application of fluorescent Lifeact fusion constructs for live-cell imaging of F-actin in N. crassa. We constructed two synthetic genes consisting of Lifeact fused to “synthetic” green fluorescent protein (sGFP) (S65T) (henceforth termed GFP) (12) or red fluorescent protein (TagRFP) (33) and expressed these constructs in various N. crassa strains. In all strain backgrounds, fluorescent Lifeact constructs clearly labeled actin patches, cables, and rings and revealed a direct association of F-actin structures with sites of cell polarization and active tip growth. Our results demonstrate the efficacy of Lifeact as a nontoxic live-cell imaging probe in N. crassa.     

Identification of a Short Spir Interaction Sequence at the C-terminal End of Formin Subgroup Proteins

The actin nucleation factors Spire and Cappuccino interact with each other and regulate essential cellular events during Drosophila oogenesis in a cooperative fashion. The interaction blocks formin actin nucleation activity and enhances the Spire activity. Analogous to Spire and Cappuccino, the mammalian homologs Spir-1 and formin-2 show a regulatory interaction. To get an understanding of the nature of the Spir-formin cooperation, we have analyzed the interaction biochemically and biophysically. Our data shows that the association of Spir-1 and formin-2 is not significantly mediated by binding of the Spir-1-KIND domain to the formin FH2 core domain. Instead, a short sequence motif C-terminal adjacent to the formin-2-FH2 domain could be characterized that mediates the interaction and is conserved among the members of the Fmn subgroup of formins. In line with this, we found that both mammalian Spir proteins, Spir-1 and Spir-2, interact with mammalian Fmn subgroup proteins formin-1 and formin-2.Basic cell biological functions such as proliferation, migration, division, and vesicle transport rely on the organization of the actin cytoskeleton. The initiation of actin polymerization from free actin monomers is regulated by actin nucleation factors (NF),² which help to overcome the kinetic barrier of spontaneous G-actin nucleation and, thus, catalyze the formation of filamentous actin structures and networks (1). To date, three different classes of NFs are described, the ARP2/3 complex, FH2 domain containing NFs of the formin superfamily, and NFs containing one or multiple WH2 domains (Spire/Cordon-bleu/Leiomodin) (2). The formin superfamily is subdivided into seven subfamilies (Dia, FRL, DAAM, Delphilin, INF, FHOD, Fmn) (3). The mechanisms of actin nucleation as well as the regulation of the NFs vary significantly between the three classes (and also show variances in between the distinct superfamilies). Spire and Cappuccino are NFs that belong to the Spire subfamily of WH2 containing nucleators and to the Fmn subfamily of the FH2 domain containing formins, respectively. In contrast to the Arp2/3 complex that nucleates branched filaments, Spire and the formin Cappuccino nucleate unbranched actin filaments (4).Almost two decades ago it was found that mutants of the two Drosophila NFs (Spire/Cappuccino) have an identical phenotype in early Drosophila oogenesis, i.e. both induce premature ooplasmic streaming (5, 6). Later it was shown that both proteins cooperate in the generation of a dynamic actin mesh in the oocyte that prevents premature ooplasmic streaming (7). Spire and Cappuccino do not solely have the same mutant phenotype; the proteins also physically interact and cross-regulate each other. The Cappuccino C-terminal half, encoding the FH2 domain and flanking sequences, enhances the nucleation activity of Spire, whereas the nucleation activity of Cappuccino is decreased in the presence of the Spire-KIND domain (8).Cappuccino belongs to the Fmn subgroup of formins (3, 9). In mammals, two Fmn subgroup members (formin-1, formin-2) and two Spir proteins (Spir-1, Spir-2) exist (3, 10). The formin-2 and spir-1 genes are coexpressed in the developing and adult nervous system, and the proteins interact analogous to their Drosophila counterparts Spire and Cappuccino (8, 10). Several reports showed the importance of formin-2 in mouse oogenesis and here especially in the positioning of the meiotic spindle (11–14). Recently it was found that a dynamic actin mesh, as during Drosophila oogenesis, is also required for mouse oogenesis (11, 14). The correct localization of the meiotic spindle during mouse oogenesis and the resulting asymmetric division depends on an actin mesh that is built up by formin-2. Myosin-2 generates the pulling forces required for spindle movement (14). Beside the evolutionary conserved roles for the formins Cappuccino and formin-2, Spire family proteins also seem to be evolutionary conserved regulators of oocyte development. Spire genes of the African clawed frog Xenopus (pEg6) and the sea squirt Ciona savignyi (Pem-5) have been identified as maternal genes in the oocyte in analogy to its Drosophila homolog and are proposed to function in polarity during early embryogenesis (15, 16).In an initial characterization it was found that the KIND domains of Spir-1/dSpire interact with the C-terminal sequences of formin-2/Cappuccino, which encode the FH2 domains and flanking sequences (8). To gain a further understanding of the interaction and cross-regulation of the two proteins, we investigated this interaction in detail. The objective of the study was the dissection of the formin-2/Spir-1 interaction and the determination of the structural elements that are responsible for the binding. The dissection revealed a high affinity Spir-1 interaction site of formin-2, which could be mapped to the very C terminus of formin-2 adjacent to its core FH2 domain (formin Spir interaction (FSI) sequence). The FSI sequence is conserved among the members of the Fmn subgroup. Consistently we found that all mammalian members of the two distinct nucleator families, Spir-1/2 and Fmn-1/2, interact with each other.     

Dynamin2 GTPase and Cortactin Remodel Actin Filaments

The large GTPase dynamin, best known for its activities that remodel membranes during endocytosis, also regulates F-actin-rich structures, including podosomes, phagocytic cups, actin comet tails, subcortical ruffles, and stress fibers. The mechanisms by which dynamin regulates actin filaments are not known, but an emerging view is that dynamin influences F-actin via its interactions with proteins that interact directly or indirectly with actin filaments. We show here that dynamin2 GTPase activity remodels actin filaments in vitro via a mechanism that depends on the binding partner and F-actin-binding protein, cortactin. Tightly associated actin filaments cross-linked by dynamin2 and cortactin became loosely associated after GTP addition when viewed by transmission electron microscopy. Actin filaments were dynamically unraveled and fragmented after GTP addition when viewed in real time using total internal reflection fluorescence microscopy. Cortactin stimulated the intrinsic GTPase activity of dynamin2 and maintained a stable link between actin filaments and dynamin2, even in the presence of GTP. Filaments remodeled by dynamin2 GTPase in vitro exhibit enhanced sensitivity to severing by the actin depolymerizing factor, cofilin, suggesting that GTPase-dependent remodeling influences the interactions of actin regulatory proteins and F-actin. The global organization of the actomyosin cytoskeleton was perturbed in U2-OS cells depleted of dynamin2, implicating dynamin2 in remodeling actin filaments that comprise supramolecular F-actin arrays in vivo. We conclude that dynamin2 GTPase remodels actin filaments and plays a role in orchestrating the global actomyosin cytoskeleton.Controlled assembly and disassembly of actin filaments underlies movement, shape, division, trafficking of lipids and proteins of the cell and pathogenesis by infectious bacteria and viruses. Several proteins and signaling circuits modulate actin filament dynamics, including proteins that nucleate formation of new filaments, filament cross-linking proteins that stabilize branched and bundled filament arrays, and depolymerizing factors that promote filament disassembly (1). Studies with reconstituted systems show that a single actin nucleating factor, such as the Arp2/3 complex together with a nucleation-promoting factor, a barbed end capping protein to preserve the actin monomer pool and promote nucleation, and a filament disassembly factor, such as ADF/cofilin, are sufficient to establish a dynamic dendritic actin network in vitro that mimics many properties of actin networks at the leading edge of migrating cells (2–4). However, the mechanisms for coordinating the organization and dynamics of actin filaments associated with higher-order cellular structures such as the subcortical F-actin network, F-actin at focal adhesions, and actomyosin arrays are not as well understood.Considerable evidence indicates that the large GTPase dynamin, a key mediator of membrane remodeling and fission, also influences actin filaments (reviewed in Refs. 5–7). Although the mechanisms are unknown, dynamin could influence actin filaments via its interactions with a number of proteins that directly or indirectly regulate actin filament assembly, filament stability, or filament organization. For example, several protein scaffolds biochemically link dynamin and the Arp2/3 complex activating factor, N-WASP, suggesting that the machinery for de novo actin assembly may be targeted or activated by dynamin (6, 8, 9). Dynamin2 is associated with several dynamic F-actin-containing structures in vivo, including podosomes, F-actin comet tails, phagocytic cups, dynamic cortical ruffles, and pedestal structures elaborated by enteropathogenic Escherichia coli (10–20). Cortactin, which directly binds both dynamin and actin filaments, is associated with many of the same dynamic actin structures as dynamin (5, 7) and is required for both clathrin-dependent and -independent endocytosis (21, 22). Thus, dynamin-cortactin interaction may be an important link between actin filaments and dynamin during formation or turnover of F-actin-rich structures.Considerable evidence supports the notion that GTP hydrolysis by dynamin catalyzes membrane fission activity via GTPase-dependent changes in conformation (23, 24) or via GTPase-dependent cycles of assembly and disassembly (25, 26). We hypothesize that GTPase-dependent changes in dynamin linked via its interacting proteins to actin filaments or actin regulators could similarly influence actin filaments. Overexpressed, dominant negative dynamin mutant proteins impaired in binding or hydrolyzing GTP (most often the dynamin-K44A mutation) perturb a variety of F-actin-rich cellular structures, including stress fibers and focal adhesions (27, 28), dendritic spines of neurons (29), podosomes (12, 30), actin comet tails (13, 14), phagocytic cups and bacteria-induced pedestal structures (16, 19), and dynamic cortical ruffles (15, 17). In addition, F-actin of stress fibers and overall cell morphology were perturbed in Clone9 cells expressing a mutant dynamin2 protein lacking the C-terminal proline-rich domain, the domain through which dynamin2 interacts with actin regulatory factors (11). Whereas existing data indicates that the specific effects of dynamin GTPase activity on F-actin structures are cell type- and structure-specific, a general conclusion is that dynamin GTPase activity influences the organization or turnover of a subset of actin filaments.To determine the mechanisms by which dynamin2 GTPase activity influences actin filaments, we developed biochemical and microscopic approaches to quantitatively assess and observe GTPase-dependent effects on actin filaments formed in vitro with Arp2/3 complex, cortactin, and dynamin2. The activities of dynamin2 on actin filaments in vivo were examined in cells with disrupted dynamin2 function using siRNA²-mediated suppression or pharmacologic inhibition. We report that dynamin2 GTPase, together with cortactin, functions as a dynamic actin filament remodeling complex that influences the global organization of the actomyosin cytoskeleton.     

A Small Conserved Domain in the Yeast Spa2p Is Necessary and Sufficient for Its Polarized Localization

SPA2 encodes a yeast protein that is one of the first proteins to localize to sites of polarized growth, such as the shmoo tip and the incipient bud. The dynamics and requirements for Spa2p localization in living cells are examined using Spa2p green fluorescent protein fusions. Spa2p localizes to one edge of unbudded cells and subsequently is observable in the bud tip. Finally, during cytokinesis Spa2p is present as a ring at the mother–daughter bud neck. The bud emergence mutants bem1 and bem2 and mutants defective in the septins do not affect Spa2p localization to the bud tip. Strikingly, a small domain of Spa2p comprised of 150 amino acids is necessary and sufficient for localization to sites of polarized growth. This localization domain and the amino terminus of Spa2p are essential for its function in mating. Searching the yeast genome database revealed a previously uncharacterized protein which we name, Sph1p (Spa2p homolog), with significant homology to the localization domain and amino terminus of Spa2p. This protein also localizes to sites of polarized growth in budding and mating cells. SPH1, which is similar to SPA2, is required for bipolar budding and plays a role in shmoo formation. Overexpression of either Spa2p or Sph1p can block the localization of either protein fused to green fluorescent protein, suggesting that both Spa2p and Sph1p bind to and are localized by the same component. The identification of a 150–amino acid domain necessary and sufficient for localization of Spa2p to sites of polarized growth and the existence of this domain in another yeast protein Sph1p suggest that the early localization of these proteins may be mediated by a receptor that recognizes this small domain.Polarized cell growth and division are essential cellular processes that play a crucial role in the development of eukaryotic organisms. Cell fate can be determined by cell asymmetry during cell division (Horvitz and Herskowitz, 1992; Cohen and Hyman, 1994; Rhyu and Knoblich, 1995). Consequently, the molecules involved in the generation and maintenance of cell asymmetry are important in the process of cell fate determination. Polarized growth can occur in response to external signals such as growth towards a nutrient (Rodriguez-Boulan and Nelson, 1989; Eaton and Simons, 1995) or hormone (Jackson and Hartwell, 1990a , b ; Segall, 1993; Keynes and Cook, 1995) and in response to internal signals as in Caenorhabditis elegans (Goldstein et al., 1993; Kimble, 1994; Priess, 1994) and Drosophila melanogaster (St Johnston and Nusslein-Volhard, 1992; Anderson, 1995) early development. Saccharomyces cerevisiae undergo polarized growth towards an external cue during mating and to an internal cue during budding. Polarization towards a mating partner (shmoo formation) and towards a new bud site requires a number of proteins (Chenevert, 1994; Chant, 1996; Drubin and Nelson, 1996). Many of these proteins are necessary for both processes and are localized to sites of polarized growth, identified by the insertion of new cell wall material (Tkacz and Lampen, 1972; Farkas et al., 1974; Lew and Reed, 1993) to the shmoo tip, bud tip, and mother–daughter bud neck. In yeast, proteins localized to growth sites include cytoskeletal proteins (Adams and Pringle, 1984; Kilmartin and Adams, 1984; Ford, S.K., and J.R. Pringle. 1986. Yeast. 2:S114; Drubin et al., 1988; Snyder, 1989; Snyder et al., 1991; Amatruda and Cooper, 1992; Lew and Reed, 1993; Waddle et al., 1996), neck filament components (septins) (Byers and Goetsch, 1976; Kim et al., 1991; Ford and Pringle, 1991; Haarer and Pringle, 1987; Longtine et al., 1996), motor proteins (Lillie and Brown, 1994), G-proteins (Ziman, 1993; Yamochi et al., 1994; Qadota et al., 1996), and two membrane proteins (Halme et al., 1996; Roemer et al., 1996; Qadota et al., 1996). Septins, actin, and actin-associated proteins localize early in the cell cycle, before a bud or shmoo tip is recognizable. How this group of proteins is localized to and maintained at sites of cell growth remains unclear.Spa2p is one of the first proteins involved in bud formation to localize to the incipient bud site before a bud is recognizable (Snyder, 1989; Snyder et al., 1991; Chant, 1996). Spa2p has been localized to where a new bud will form at approximately the same time as actin patches concentrate at this region (Snyder et al., 1991). An understanding of how Spa2p localizes to incipient bud sites will shed light on the very early stages of cell polarization. Later in the cell cycle, Spa2p is also found at the mother–daughter bud neck in cells undergoing cytokinesis. Spa2p, a nonessential protein, has been shown to be involved in bud site selection (Snyder, 1989; Zahner et al., 1996), shmoo formation (Gehrung and Snyder, 1990), and mating (Gehrung and Snyder, 1990; Chenevert et al., 1994; Yorihuzi and Ohsumi, 1994; Dorer et al., 1995). Genetic studies also suggest that Spa2p has a role in cytokinesis (Flescher et al., 1993), yet little is known about how this protein is localized to sites of polarized growth.We have used Spa2p green fluorescent protein (GFP)¹ fusions to investigate the early localization of Spa2p to sites of polarized growth in living cells. Our results demonstrate that a small domain of ∼150 amino acids of this large 1,466-residue protein is sufficient for targeting to sites of polarized growth and is necessary for Spa2p function. Furthermore, we have identified and characterized a novel yeast protein, Sph1p, which has homology to both the Spa2p amino terminus and the Spa2p localization domain. Sph1p localizes to similar regions of polarized growth and sph1 mutants have similar phenotypes as spa2 mutants.     

Global Analysis of Cdc14 Dephosphorylation Sites Reveals Essential Regulatory Role in Mitosis and Cytokinesis

Degradation of the M phase cyclins triggers the exit from M phase. Cdc14 is the major phosphatase required for the exit from the M phase. One of the functions of Cdc14 is to dephosphorylate and activate the Cdh1/APC/C complex, resulting in the degradation of the M phase cyclins. However, other crucial targets of Cdc14 for mitosis and cytokinesis remain to be elucidated. Here we systematically analyzed the positions of dephosphorylation sites for Cdc14 in the budding yeast Saccharomyces cerevisiae. Quantitative mass spectrometry identified a total of 835 dephosphorylation sites on 455 potential Cdc14 substrates in vivo. We validated two events, and through functional studies we discovered that Cdc14-mediated dephosphorylation of Smc4 and Bud3 is essential for proper mitosis and cytokinesis, respectively. These results provide insight into the Cdc14-mediated pathways for exiting the M phase.All cells proliferate following a fixed, highly coordinated cycle. Mitosis especially requires elaborate coordination for proper chromosome segregation, mitotic spindle disassembly, and cytokinesis. Much of this activity is facilitated by numerous, diverse phosphorylation and dephosphorylation signals that orchestrate the precise progression of M phase.Prior to mitosis, sister chromatids resulting from DNA replication during S phase are held together by the cohesion complex. Then, during prophase, chromosomes are condensed by the condensin (Smc2/4) complex (1) and microtubules are remodeled to form the mitotic spindle (2). Subsequently, in metaphase, the microtubules of the spindle apparatus attach to the chromosome kinetochores (3) and dissolution of the sister chromatids is triggered by the separase-mediated cleavage of cohesin (4, 5). Finally, Cdc14, Cdh1, and APC/C work together in telophase to degrade the M phase cyclins (6), promote decondensation of chromosomes (7), and finish cytokinesis (8, 9).Cdc14, a dual-specificity phosphatase that removes the phosphate group on both phosphotyrosine and phosphoserine/threonine residues (10), is required for mitosis (11, 12). Specifically, Cdc14 function is essential in late M phase: cells carrying a defective mutation arrest in telophase (13), whereas overexpression of Cdc14 results in G1 arrest (12). Cdc14 triggers mitotic cyclin-dependent kinase (CDK)¹ inactivation, enabling cells to exit mitosis through dephosphorylation and activation of the inhibitors of CDKs. At interphase, Cdc14 is a subunit of the mitotic exit network (14–17), which usually localizes to the nucleolus. However, the Cdc14 early anaphase release network initiates the release of Cdc14 from its inhibitor, Net1/Cfi1 (18), and the mitotic exit network promotes further release of Cdc14 from its inhibitor, allowing it to spread into the nucleus and cytoplasm, where it dephosphorylates its major targets (8, 9), leading to exit from mitosis. In addition to this essential role in late M phrase, Cdc14 substrates have also been identified in other stages of the cell cycle (19).Cdc14 putatively regulates 27 proteins (19–22). Some studies have documented the substrates of Cdc14 via in vitro phosphatase assay, whereas others have provided in vivo evidence. However, dephosphorylation sites have been identified for only five of the target proteins (17, 22–25), suggesting that spurious relationships cannot be ruled out. Also, experiments have not been carried out to demonstrate whether these modifications entail direct or indirect regulation. Therefore, our understanding of Cdc14 function and regulation during mitosis in metazoans is incomplete. Conceivably, Cdc14 may regulate many more substrates involved in aspects of chromosome condensation and cytokinesis. To examine this possibility we performed a systematic phosphoproteomic screen to identify new in vivo pathways regulated by Cdc14. Using this approach, we identified both known and potentially novel substrates of Cdc14, as well as their dephosphorylation sites. Many potentially novel substrates are physically associated with Cdc14 in public databases. We also provide biochemical evidence for direct dephosphorylation of the substrates, characterize the specificity of dephosphorylation in two substrates, Smc4 and Bud3, and further study their regulation and critical role in mitosis and cytokinesis.     

Vinculin Nucleates Actin Polymerization and Modifies Actin Filament Structure

Vinculin links integrins to the actin cytoskeleton by binding F-actin. Little is known with respect to how this interaction occurs or affects actin dynamics. Here we assess the consequence of the vinculin tail (VT) on actin dynamics by examining its binding to monomeric and filamentous yeast actins. VT causes pyrene-labeled G-actin to polymerize in low ionic strength buffer (G-buffer), conditions that normally do not promote actin polymerization. Analysis by electron microscopy shows that, under these conditions, the filaments form small bundles at low VT concentrations, which gradually increase in size until saturation occurs at a ratio of 2 VT:1 actin. Addition of VT to pyrene-labeled mutant yeast G-actin (S265C) produced a fluorescence excimer band, which requires a relatively normal filament geometry. In higher ionic strength polymerization-promoting F-buffer, substoichiometric amounts of VT accelerate the polymerization of pyrene-labeled WT actin. However, the amplitude of the pyrene fluorescence caused by actin polymerization is quenched as the VT concentration increases without an effect on net actin polymerization as determined by centrifugation assays. Finally, addition of VT to preformed pyrene-labeled S265C F-actin causes a concentration-dependent decrease in the maximum amplitude of the pyrene fluorescence band demonstrating the ability of VT to remodel the conformation of the actin filament. These observations support the idea that vinculin can link adhesion plaques to the cytoskeleton by initiating the formation of bundled actin filaments or by remodeling existing filaments.Cell migration is critical for embryonic development, adult homeostasis, inflammatory responses, and wound healing. To migrate, a cell must coordinate a number of different inputs into appropriate cellular responses. The cell must polarize in the direction of migration and extend lamellipodial and/or filopodial protrusions. Nascent adhesions that assemble within the branched actin network of the lamellipodium must link to the underlying actin cytoskeleton. This process allows for the maturation of adhesions to structures that anchor the protrusion. These adhesions also provide the traction forces necessary to pull the cell body forward and break older adhesions at the cell rear. Perturbation of any of these events affects a cell''s migratory ability. For example, nascent adhesions that do not form linkages to the actin cytoskeleton cannot effectively anchor the protrusion to the substratum. The result is an extension that folds back upon itself, forming a membrane ruffle that cannot provide the traction forces necessary for migration.How adhesions establish links to the underlying actin cytoskeleton has been an area of intense investigation. Integrin-containing structures are active areas of actin polymerization suggesting that adhesion plaques can initiate actin filament formation (reviewed in Refs. 1–3). Focal complexes are small integrin clusters that are found exclusively at the tips of lamellipodia and filopodia. Formation of these structures is closely coupled with actin assembly in protruding regions of cells. Accumulating evidence indicates that adhesion complex components recruit the Arp2/3 complex, a potent nucleator of actin polymerization. Our work (4) and that of others (5–7) demonstrates that the Arp2/3 complex is recruited to focal complexes or transient adhesion structures reminiscent of focal complexes by binding vinculin. FAK has also been implicated in linking focal complexes to the actin cytoskeleton by virtue of its ability to recruit and activate the Arp2/3 complex (8). Furthermore, efficient focal complex assembly requires the actin-binding protein, cortactin, which could affect adhesion assembly by interacting with the Arp2/3 complex (9). Hence, many of the known mechanisms for initiating filament formation involve recruitment of the Arp2/3 complex, which initiates the formation of branched actin filaments (55). It is surprising then that the earliest detectable forms of actin-associated adhesions are interconnected by short actin bundles, not branched filaments (10). These observations suggest that our current understanding for how nascent adhesions initiate filament formation is incomplete.The earliest detectable actin-associated adhesions are “dots or doublets of dots” and are highly enriched in integrins, paxillin, and vinculin (10), suggesting that one of these molecules has the capability to initiate actin filament formation from such a plaque. Vinculin has long been implicated in linking adhesion plaques to the actin cytoskeleton by virtue of the ability of its tail to bind (11) and bundle F-actin (12). The interaction of vinculin with actin has been extensively studied from the perspective of vinculin (11, 13–23). Studies of recombinant proteins identified two regions of the vinculin tail (VT)² that bind F-actin independently (21, 17), but mapping these sites onto the VT crystal structure reveals that these peptides do not correspond to distinct sites (25). Upon binding actin, vinculin undergoes a conformational change that promotes dimerization suggesting that vinculin self-association may be important for its bundling activities (15).Less is known with respect to the effect of vinculin on actin filament formation and structure. This lack of knowledge stems from the fact that many of the early studies showed vinculin to have no effect on actin dynamics (26–28). However, these experiments were performed using chicken gizzard vinculin, which exists almost exclusively in a conformation where the actin binding sites are inaccessible, or from preparations that contain contaminants that produce false negatives (29). More recently, recombinant VT proteins were shown to cross-link and bundle actin (23). However, the interaction of vinculin with G-actin and the effect of vinculin on actin filament dynamics have not been explored. In this study, we have assessed the interaction of vinculin with pyrene-labeled wild-type and mutant yeast actins. We show that the VT can promote the formation of an actin nucleus from which filaments arise and alter the assembly and structure of actin filaments. These findings provide novel insights into how adhesion plaques may be linked to the actin cytoskeleton.     

Analysis of Free Energy Signals Arising from Nucleotide Hybridization Between rRNA and mRNA Sequences during Translation in Eubacteria

A decoding algorithm is tested that mechanistically models the progressive alignments that arise as the mRNA moves past the rRNA tail during translation elongation. Each of these alignments provides an opportunity for hybridization between the single-stranded, -terminal nucleotides of the 16S rRNA and the spatially accessible window of mRNA sequence, from which a free energy value can be calculated. Using this algorithm we show that a periodic, energetic pattern of frequency 1/3 is revealed. This periodic signal exists in the majority of coding regions of eubacterial genes, but not in the non-coding regions encoding the 16S and 23S rRNAs. Signal analysis reveals that the population of coding regions of each bacterial species has a mean phase that is correlated in a statistically significant way with species () content. These results suggest that the periodic signal could function as a synchronization signal for the maintenance of reading frame and that codon usage provides a mechanism for manipulation of signal phase.[1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32]     

Proinsulin and the Genetics of Diabetes Mellitus

Insulin plays a central role in the regulation of vertebrate metabolism. The hormone, the post-translational product of a single-chain precursor, is a globular protein containing two chains, A (21 residues) and B (30 residues). Recent advances in human genetics have identified dominant mutations in the insulin gene causing permanent neonatal-onset DM² (1–4). The mutations are predicted to block folding of the precursor in the ER of pancreatic β-cells. Although expression of the wild-type allele would in other circumstances be sufficient to maintain homeostasis, studies of a corresponding mouse model (5–7) suggest that the misfolded variant perturbs wild-type biosynthesis (8, 9). Impaired β-cell secretion is associated with ER stress, distorted organelle architecture, and cell death (10). These findings have renewed interest in insulin biosynthesis (11–13) and the structural basis of disulfide pairing (14–19). Protein evolution is constrained not only by structure and function but also by susceptibility to toxic misfolding.Insulin plays a central role in the regulation of vertebrate metabolism. The hormone, the post-translational product of a single-chain precursor, is a globular protein containing two chains, A (21 residues) and B (30 residues). Recent advances in human genetics have identified dominant mutations in the insulin gene causing permanent neonatal-onset DM² (1–4). The mutations are predicted to block folding of the precursor in the ER of pancreatic β-cells. Although expression of the wild-type allele would in other circumstances be sufficient to maintain homeostasis, studies of a corresponding mouse model (5–7) suggest that the misfolded variant perturbs wild-type biosynthesis (8, 9). Impaired β-cell secretion is associated with ER stress, distorted organelle architecture, and cell death (10). These findings have renewed interest in insulin biosynthesis (11–13) and the structural basis of disulfide pairing (14–19). Protein evolution is constrained not only by structure and function but also by susceptibility to toxic misfolding.     

Algorithms for Finding Small Attractors in Boolean Networks

A Boolean network is a model used to study the interactions between different genes in genetic regulatory networks. In this paper, we present several algorithms using gene ordering and feedback vertex sets to identify singleton attractors and small attractors in Boolean networks. We analyze the average case time complexities of some of the proposed algorithms. For instance, it is shown that the outdegree-based ordering algorithm for finding singleton attractors works in time for , which is much faster than the naive time algorithm, where is the number of genes and is the maximum indegree. We performed extensive computational experiments on these algorithms, which resulted in good agreement with theoretical results. In contrast, we give a simple and complete proof for showing that finding an attractor with the shortest period is NP-hard.[1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32]     

Phosphorylation Dependence of Hsp27 Multimeric Size and Molecular Chaperone Function

The molecular chaperone Hsp27 exists as a distribution of large oligomers that are disassembled by phosphorylation at Ser-15, -78, and -82. It is controversial whether the unphosphorylated Hsp27 or the widely used triple Ser-to-Asp phospho-mimic mutant is the more active molecular chaperone in vitro. This question was investigated here by correlating chaperone activity, as measured by the aggregation of reduced insulin or α-lactalbumin, with Hsp27 self-association as monitored by analytical ultracentrifugation. Furthermore, because the phospho-mimic is generally assumed to reproduce the phosphorylated molecule, the size and chaperone activity of phosphorylated Hsp27 were compared with that of the phospho-mimic. Hsp27 was triply phosphorylated by MAPKAP-2 kinase, and phosphorylation was tracked by urea-PAGE. An increasing degree of suppression of insulin or α-lactalbumin aggregation correlated with a decreasing Hsp27 self-association, which was the least for phosphorylated Hsp27 followed by the mimic followed by the unphosphorylated protein. It was also found that Hsp27 added to pre-aggregated insulin did not reverse aggregation but did inhibit these aggregates from assembling into even larger aggregates. This chaperone activity appears to be independent of Hsp27 phosphorylation. In conclusion, the most active chaperone of insulin and α-lactalbumin was the Hsp27 (elongated) dimer, the smallest Hsp27 subunit observed under physiological conditions. Next, the Hsp27 phospho-mimic is only a partial mimic of phosphorylated Hsp27, both in self-association and in chaperone function. Finally, the efficient inhibition of insulin aggregation by Hsp27 dimer led to the proposal of two models for this chaperone activity.Oligomeric heat shock protein 27 (Hsp27)² is a ubiquitous mammalian protein with a variety of functions in health and disease (1–8). These functions include ATP-independent chaperone activity in response to environmental stress, e.g. heat shock and oxidative stress, control of apoptosis, and regulation of actin cytoskeleton dynamics. Hsp27 is a member of the α-crystallin small heat shock protein family of which αB-crystallin is the archetype. These proteins are characterized by an α-crystallin domain of 80–90 residues consisting of roughly eight β-strands that form an intermolecular β-sheet interaction interface within a dimer, the basic building subunit of the oligomer (2, 4, 9–11).Hsp27 is in equilibrium between high molecular weight oligomers and much lower molecular weight multimers. It has been reported that unphosphorylated Hsp27 includes predominantly a distribution of high molecular species ranging in size from 12-mer to 35-mer (12–19). Phosphorylation of Hsp27 at serines 15, 78, and 82 by the p38-activated MAPKAP-2 kinase (20–22) or the use of the triple Ser-to-Asp phospho-mimic results in a major shift in the equilibrium toward much smaller multimers (23) and in an alteration of its function (1, 3, 6, 7, 24, 25). The size distribution of the smaller species has been reported to be between monomer and tetramer (12–16, 18, 19).Small heat shock proteins, including Hsp27, behave as ATP-independent molecular chaperones during cellular heat shock. They bind partially unfolded proteins and prevent their aggregation until the proteins can be refolded by larger ATP-dependent chaperones or are digested (7, 8, 26). This function includes the up-regulation and/or phosphorylation of Hsp27.It is not entirely clear what the role of Hsp27 size and phosphorylation state plays in its heat shock function because there are conflicting results in the literature. Some in vitro studies concluded that the unphosphorylated oligomeric Hsp27 (or the murine isoform Hsp25) protects proteins against aggregation better than does the phosphorylation mimic (13, 19, 27), whereas others found no difference (16, 28, 29), and still other studies found that the mimic protects better than does the unphosphorylated wild type (27, 30, 31). In-cell studies found that phosphorylation of Hsp27 was essential for thermo-protection of actin filaments (32), and the Hsp27 phosphorylation mimic decreased inclusion body formation better than did unphosphorylated Hsp27 (33). This study was undertaken to investigate the molecular chaperone function of Hsp27 by correlating chaperone activity with Hsp27 size and by comparing fully phosphorylated Hsp27 with its phospho-mimic.     

Splitting the BLOSUM Score into Numbers of Biological Significance

Mathematical tools developed in the context of Shannon information theory were used to analyze the meaning of the BLOSUM score, which was split into three components termed as the BLOSUM spectrum (or BLOSpectrum). These relate respectively to the sequence convergence (the stochastic similarity of the two protein sequences), to the background frequency divergence (typicality of the amino acid probability distribution in each sequence), and to the target frequency divergence (compliance of the amino acid variations between the two sequences to the protein model implicit in the BLOCKS database). This treatment sharpens the protein sequence comparison, providing a rationale for the biological significance of the obtained score, and helps to identify weakly related sequences. Moreover, the BLOSpectrum can guide the choice of the most appropriate scoring matrix, tailoring it to the evolutionary divergence associated with the two sequences, or indicate if a compositionally adjusted matrix could perform better.[1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29]     

Myosin II Tailpiece Determines Its Paracrystal Structure, Filament Assembly Properties, and Cellular Localization

Non muscle myosin II (NMII) is a major motor protein present in all cell types. The three known vertebrate NMII isoforms share high sequence homology but play different cellular roles. The main difference in sequence resides in the C-terminal non-helical tailpiece (tailpiece). In this study we demonstrate that the tailpiece is crucial for proper filament size, overcoming the intrinsic properties of the coiled-coil rod. Furthermore, we show that the tailpiece by itself determines the NMII filament structure in an isoform-specific manner, thus providing a possible mechanism by which each NMII isoform carries out its unique cellular functions. We further show that the tailpiece determines the cellular localization of NMII-A and NMII-B and is important for NMII-C role in focal adhesion complexes. We mapped NMII-C sites phosphorylated by protein kinase C and casein kinase II and showed that these phosphorylations affect its solubility properties and cellular localization. Thus phosphorylation fine-tunes the tailpiece effects on the coiled-coil rod, enabling dynamic regulation of NMII-C assembly. We thus show that the small tailpiece of NMII is a distinct domain playing a role in isoform-specific filament assembly and cellular functions.Non muscle myosin II (NMII)² is a major motor protein present in all cell types participating in crucial processes, including cytokinesis, surface attachment, and cell movement (1–3). NMII units are hexamers of two long heavy chains with two pairs of light chains attached. NMII heavy chain is composed of a globular head containing the actin binding and force generating ATPase domains, followed by a large coiled-coil rod that terminates with a short non-helical tailpiece (tailpiece). To carry out its cellular functions, NMII assembles into dimers and higher order filaments by interactions of the coiled-coil rod (4). The assembly process is governed by electrostatic interactions between adjacent coiled-coil rods containing alternating charged regions with specific periodicity (5–9) and is enhanced by activation of the motor domain through regulatory light chain phosphorylation (10–12). The charge periodicity also determines the register and orientation of each NMII hexamer in the filament. Additionally the C-terminal region of the coiled-coil rod contains a distinctive positively charged region and the assembly-competence domains that are crucial for proper filament assembly (5–9, 13).Three isoforms of NMII (termed NMII-A, NMII-B, and NMII-C) have been identified in mammals (14–16). Although NMII isoforms share somewhat overlapping roles, each isoform has distinctive tissue distribution and specific functions. NMII-A is important for neural growth cone retraction (17, 18) and is distributed to the front of migrating endothelial cells (19). While NMII-B participates in growth cone advancement (20) and was detected in the retracting tails of migrating endothelial cells (19). Furthermore NMII-A and NMII-B have an opposing effect on motility, since depletion of NMII-A leads to increased motility while NMII-B depletion hinders motility (21, 22). NMII-C plays a role in cytokinesis (23) and has distinct distribution in neuronal cells (24). Furthermore one NMII isoform only partly rescue cells in which siRNA was used to reduce the expression of another isoform (23, 25). This functional diversity is achieved despite a significant amino acid sequence identity between the isoforms (overall 64–80%), and the origin of these differential distributions and functions is not completely understood.Recent studies suggest that the C-terminal portion of NMII-A and NMII-B, particularly the last ∼170 amino acids, is responsible for the differential distribution of these NMII isoforms (26, 27). It was shown that swapping this region between NMII-A and NMII-B resulted in chimeric proteins, which adopted cellular localization according to the C-terminal part (26). This C-terminal ∼170 amino acid coiled-coil region contains the assembly-competence domains and other regions that are critical for filament assembly (5–9, 13) as well as the non-helical tailpiece. As the small tailpiece is also an important regulator of NMII filament assembly (27, 28) capable of changing NMII filament assembly properties; and phosphorylation of NMII tailpiece was shown to interfere with filament assembly (29–33) the tailpiece may be important for allowing NMII to perform its dynamic tasks. Because the coiled-coil regions are highly conserved between NMII isoforms, while the tailpiece is the most divergent, it is therefore a good candidate for mediating NMII isoform-specific functions. However, the exact mechanism by which the tailpiece affects NMII function is not fully understood. Here we show that the tailpiece serves as an isoform-specific control mechanism modulating filament order, assembly, and cellular function.     

B Cell Receptor-induced Phosphorylation of Pyk2 and Focal Adhesion Kinase Involves Integrins and the Rap GTPases and Is Required for B Cell Spreading

Signaling by the B cell receptor (BCR) promotes integrin-mediated adhesion and cytoskeletal reorganization. This results in B cell spreading, which enhances the ability of B cells to bind antigens and become activated. Proline-rich tyrosine kinase (Pyk2) and focal adhesion kinase (FAK) are related cytoplasmic tyrosine kinases that regulate cell adhesion, cell morphology, and cell migration. In this report we show that BCR signaling and integrin signaling collaborate to induce the phosphorylation of Pyk2 and FAK on key tyrosine residues, a modification that increases the kinase activity of Pyk2 and FAK. Activation of the Rap GTPases is critical for BCR-induced integrin activation as well as for BCR- and integrin-induced reorganization of the actin cytoskeleton. We now show that Rap activation is essential for BCR-induced phosphorylation of Pyk2 and for integrin-induced phosphorylation of Pyk2 and FAK. Moreover Rap-dependent phosphorylation of Pyk2 and FAK required an intact actin cytoskeleton as well as actin dynamics, suggesting that Rap regulates Pyk2 and FAK via its effects on the actin cytoskeleton. Importantly B cell spreading induced by BCR/integrin co-stimulation or by integrin engagement was inhibited by short hairpin RNA-mediated knockdown of either Pyk2 or FAK expression and by treatment with PF-431396, a chemical inhibitor that blocks the kinase activities of both Pyk2 and FAK. Thus Pyk2 and FAK are downstream targets of the Rap GTPases that play a key role in regulating B cell morphology.Antibodies (Abs)² made by B lymphocytes play a critical role in host defense against infection. Antigen-induced signaling by the B cell receptor (BCR) initiates an activation program that leads to B cell proliferation and subsequent differentiation into Ab-producing cells. BCR clustering by antigens or by anti-immunoglobulin (anti-Ig) Abs used as surrogate antigens initiates multiple signaling pathways that control gene expression, cell survival, and proliferation pathways (1–3).BCR signaling also promotes integrin activation (4, 5), localized actin polymerization, reorganization of the actin cytoskeleton, and changes in B cell morphology (6, 7), all of which may facilitate B cell activation. Integrin activation and cell spreading is critical for the activation of B cells by membrane-bound antigens. Macrophages, dendritic cells, and follicular dendritic cells can present arrays of captured antigens to B cells (8, 9), and this may be one of the main ways in which B cells encounter antigens (10). BCR-induced integrin activation prolongs the interaction between the B cell and the antigen-presenting cell and also allows the B cell to spread on the surface of the antigen-presenting cell such that more BCRs can encounter and bind membrane-bound antigens (11). Subsequent contraction of the B cell membrane allows the B cells to gather the BCR-bound antigen into an immune synapse in which clustered antigen-engaged BCRs are surrounded by a ring of ligand-bound integrins. Formation of this immune synapse reduces the amount of antigen that is required for B cell activation (12, 13).Recent work has shown that B cells in lymphoid organs may contact soluble antigens by extending membrane processes into a highly organized network of lymph-filled conduits (14). These conduits are created by fibroblastic reticular cells that partially ensheathe collagen fibrils. In addition to being rich in collagen, fibronectin, and other extracellular matrix (ECM) components, the fibroblastic reticular cells that form these conduits express high levels of intercellular adhesion molecule-1, the ligand for the α_Lβ₂ integrin (lymphocyte function-associated antigen-1 (LFA-1)) on B cells (10). Thus B cells interacting with these conduits are likely to be in contact with integrin ligands, and integrin-dependent spreading may enhance the ability of B cells to extend membrane processes into the fibroblastic reticular cell conduit.In addition to promoting cell spreading, integrins can act as co-stimulatory receptors that enhance signaling by many receptors including the T cell receptor and the BCR (15–17). Thus signaling proteins that regulate B cell spreading and that are also targets of BCR/integrin co-stimulation may play a key role in the activation of B cells by membrane-bound antigens as well as soluble antigens that are delivered to lymphoid organs by fibroblastic reticular cell conduits.Proline-rich tyrosine kinase (Pyk2) and focal adhesion kinase (FAK) are related non-receptor protein-tyrosine kinases that integrate signals from multiple receptors and play an important role in regulating cell adhesion, cell morphology, and cell migration in many cell types (18–20). Integrins, receptor tyrosine kinases, antigen receptors, and G protein-coupled chemokine receptors all stimulate tyrosine phosphorylation of Pyk2 and FAK, a modification that increases the enzymatic activity of these kinases and allows them to bind SH2 domain-containing signaling proteins (21). FAK, which is expressed in almost all tissues (21), is a focal adhesion component that mediates integrin-dependent cell migration (22), cell spreading, and cell adhesion (18) in adherent cells as well as co-clustering of LFA-1 with the T cell receptor in lymphocytes (23). Pyk2 is expressed mainly in hematopoietic cells, osteoclasts, and the central nervous system (24) and is critical for chemokine-induced migration of B cells, macrophages, and natural killer cells (20, 25, 26) as well as the spreading of osteoclasts on vitronectin (27). FAK and Pyk2 are thought to mediate overlapping but distinct functions because Pyk2 expression only partially reverses the cell adhesion and migration defects in FAK-deficient fibroblasts (28).In B cells, clustering of the BCR, β₁ integrins, or β₇ integrins induces tyrosine phosphorylation of both Pyk2 and FAK (29–33). FAK is involved in the chemokine-induced adhesion of B cell progenitors (34), and Pyk2 is required for chemokine-induced migration of mature B cells (25). However, the role of these kinases in BCR- and integrin-induced B cell spreading has not been investigated, and the signaling pathways that link the BCR and integrins to tyrosine phosphorylation of Pyk2 and FAK have not been elucidated.We have shown previously that the ability of the BCR to induce integrin activation, B cell spreading, and immune synapse formation requires activation of the Rap GTPases (6, 17). In addition to binding effector proteins such as RapL and Rap1-interacting adaptor molecule (RIAM) that promote integrin activation (35–37), the active GTP-bound forms of Rap1 and Rap2 bind multiple proteins that control actin dynamics and cell morphology (38). Moreover we showed that BCR/integrin-induced phosphorylation of Pyk2 in B cells is dependent on Rap activation (17). However, this previous study did not address how Rap-GTP links the BCR and integrins to Pyk2 phosphorylation, whether Rap activation is important for FAK phosphorylation in B cells, or whether B cell spreading is regulated by Pyk2 or FAK. We now show that Pyk2 and FAK are differentially expressed and localized in B cells, that Pyk2 and FAK are important for B cell spreading, and that integrin engagement enhances BCR-induced phosphorylation of Pyk2 and FAK, a process that depends on both Rap activation and actin dynamics.