Tyrosine Phosphorylation at a Site Highly Conserved in the L1 Family of Cell Adhesion Molecules Abolishes Ankyrin Binding and Increases Lateral Mobility of Neurofascin

This paper presents evidence that a member of the L1 family of ankyrin-binding cell adhesion molecules is a substrate for protein tyrosine kinase(s) and phosphatase(s), identifies the highly conserved FIGQY tyrosine in the cytoplasmic domain as the principal site of phosphorylation, and demonstrates that phosphorylation of the FIGQY tyrosine abolishes ankyrin-binding activity. Neurofascin expressed in neuroblastoma cells is subject to tyrosine phosphorylation after activation of tyrosine kinases by NGF or bFGF or inactivation of tyrosine phosphatases with vanadate or dephostatin. Furthermore, both neurofascin and the related molecule Nr-CAM are tyrosine phosphorylated in a developmentally regulated pattern in rat brain. The FIGQY sequence is present in the cytoplasmic domains of all members of the L1 family of neural cell adhesion molecules. Phosphorylation of the FIGQY tyrosine abolishes ankyrin binding, as determined by coimmunoprecipitation of endogenous ankyrin and in vitro ankyrin-binding assays. Measurements of fluorescence recovery after photobleaching demonstrate that phosphorylation of the FIGQY tyrosine also increases lateral mobility of neurofascin expressed in neuroblastoma cells to the same extent as removal of the cytoplasmic domain. Ankyrin binding, therefore, appears to regulate the dynamic behavior of neurofascin and is the target for regulation by tyrosine phosphorylation in response to external signals. These findings suggest that tyrosine phosphorylation at the FIGQY site represents a highly conserved mechanism, used by the entire class of L1-related cell adhesion molecules, for regulation of ankyrin-dependent connections to the spectrin skeleton.Vertebrate L1, neurofascin, neuroglial cell adhesion molecule (Ng-CAM),¹ Ng-CAM–related cell adhesion molecule (Nr-CAM), and Drosophila neuroglian are members of a family of nervous system cell adhesion molecules that possess variable extracellular domains comprised of Ig and fibronectin type III domains and a relatively conserved cytoplasmic domain (Grumet, 1991; Hortsch and Goodman, 1991; Rathgen and Jessel, 1991; Sonderegger and Rathgen, 1992; Hortsch, 1996). Members of this family, including a number of alternatively spliced forms, are abundant in the nervous system during early development as well as in adults. Neurofascin and Nr-CAM, for example, constitute ∼0.5% of the total membrane protein in adult brain (Davis et al., 1993; Davis and Bennett, 1994). Cellular functions attributed to the L1 family include axon fasciculation (Stallcup and Beasley, 1985; Landmesser et al., 1988; Brummendorf and Rathjen, 1993; Bastmeyer et al., 1995; Itoh et al., 1995; Magyar-Lehmann et al., 1995), axonal guidance (van den Pol and Kim, 1993; Liljelund et al., 1994; Brittis and Silver, 1995; Brittis et al., 1995; Lochter et al., 1995; Wong et al., 1996), neurite extension (Chang et al., 1987; Felsenfeld et al., 1994; Hankin and Lagenaur, 1994; Ignelzi et al., 1994; Williams et al., 1994a ,b,c,d; Doherty et al., 1995; Zhao and Siu, 1995), a role in long term potentiation (Luthl et al., 1994), synaptogenesis (Itoh et al., 1995), and myelination (Wood et al., 1990). The potential clinical importance of this group of proteins has been emphasized by the findings that mutations in the L1 gene on the X chromosome are responsible for developmental anomalies including hydrocephalus and mental retardation (Rosenthal et al., 1992; Jouet et al., 1994; Wong et al., 1995).The conserved cytoplasmic domains of L1 family members include a binding site for the membrane skeletal protein ankyrin. This interaction was first described for neurofascin (Davis et. al., 1993) and subsequently has been observed for L1, Nr-CAM (Davis and Bennett, 1994), and Drosophila neuroglian (Dubreuil et al., 1996). The membrane-binding domain of ankyrin contains two distinct sites for neurofascin and has the potential to promote lateral association of neurofascin and presumably other L1 family members (Michaely and Bennett, 1995). Nodes of Ranvier are physiologically relevant axonal sites where ankyrin and L1 family members collaborate, based on findings of colocalization of a specialized isoform of ankyrin with alternatively spliced forms of neurofascin and NrCAM in adults (Davis et al., 1996) as well as in early axonal developmental intermediates (Lambert, S., J. Davis, P. Michael, and V. Bennett. 1995. Mol. Biol. Cell. 6:98a).L1, after homophilic and/or heterophilic binding, participates in signal transduction pathways that ultimately are associated with neurite extension and outgrowth (Ignelzi et al., 1994; Williams et al., 1994a ,b,c,d; Doherty et al., 1995). L1 copurifies with a serine–threonine protein kinase (Sadoul et al., 1989) and is phosphorylated on a serine residue that is not conserved among other family members (Wong et al., 1996). L1 pathway(s) may also involve G proteins, calcium channels, and tyrosine phosphorylation (Williams et al., 1994a ,b,c,d; Doherty et al., 1995). After homophilic interactions, L1 directly activates a tyrosine signaling cascade after a lateral association of its ectodomain with the fibroblast growth factor receptor (Doherty et al., 1995). Antibodies against L1 have also been shown to activate protein tyrosine phosphatase activity in growth cones (Klinz et al., 1995). However, details of the downstream substrates of L1-promoted phosphorylation and dephosphorylation and possible roles of the cytoplasmic domain are not known.Tyrosine phosphorylation is well established to modulate cell–cell and cell–extracellular matrix interactions involving integrins and their associated proteins (Akiyama et al., 1994; Arroyo et al., 1994; Schlaepfer et al., 1994; Law et al., 1996) as well as the cadherins (Balsamo et al., 1996; Krypta et al., 1996; Brady-Kalnay et al., 1995; Shibamoto et al., 1995; Hoschuetzky et al., 1994; Matsuyoshi et al., 1992). For example, the adhesive functions of the calciumdependent cadherin cell adhesion molecule are mediated by a dynamic balance between tyrosine phosphorylation of β-catenin by TrkA and dephosphorylation via the LARtype protein tyrosine phosphatase (Krypta et al., 1996). In this example the regulation of binding among the structural proteins is the result of a coordination between classes of protein kinases and protein phosphatases.This study presents evidence that neurofascin, expressed in a rat neuroblastoma cell line, is a substrate for both tyrosine kinases and protein tyrosine phosphatases at a tyrosine residue conserved among all members of the L1 family. Site-specific tyrosine phosphorylation promoted by both tyrosine kinase activators (NGF and bFGF) and protein tyrosine phosphatase inhibitors (dephostatin and vanadate) is a strong negative regulator of the neurofascin– ankyrin binding interaction and modulates the membrane dynamic behavior of neurofascin. Furthermore, neurofascin and, to a lesser extent Nr-CAM, are also shown here to be tyrosine phosphorylated in developing rat brain, implying a physiological relevance to this phenomenon. These results indicate that neurofascin may be a target for the coordinate control over phosphorylation that is elicited by protein kinases and phosphatases during in vivo tyrosine phosphorylation cascades. The consequent decrease in ankyrin-binding capacity due to phosphorylation of neurofascin could represent a general mechanism among the L1 family members for regulation of membrane–cytoskeletal interactions in both developing and adult nervous systems.     

The Amino-terminal Domain of Desmoplakin Binds to Plakoglobin and Clusters Desmosomal Cadherin–Plakoglobin Complexes

The desmosome is a highly organized plasma membrane domain that couples intermediate filaments to the plasma membrane at regions of cell–cell adhesion. Desmosomes contain two classes of cadherins, desmogleins, and desmocollins, that bind to the cytoplasmic protein plakoglobin. Desmoplakin is a desmosomal component that plays a critical role in linking intermediate filament networks to the desmosomal plaque, and the amino-terminal domain of desmoplakin targets desmoplakin to the desmosome. However, the desmosomal protein(s) that bind the amino-terminal domain of desmoplakin have not been identified. To determine if the desmosomal cadherins and plakoglobin interact with the amino-terminal domain of desmoplakin, these proteins were co-expressed in L-cell fibroblasts, cells that do not normally express desmosomal components. When expressed in L-cells, the desmosomal cadherins and plakoglobin exhibited a diffuse distribution. However, in the presence of an amino-terminal desmoplakin polypeptide (DP-NTP), the desmosomal cadherins and plakoglobin were observed in punctate clusters that also contained DP-NTP. In addition, plakoglobin and DP-NTP were recruited to cell–cell interfaces in L-cells co-expressing a chimeric cadherin with the E-cadherin extracellular domain and the desmoglein-1 cytoplasmic domain, and these cells formed structures that were ultrastructurally similar to the outer plaque of the desmosome. In transient expression experiments in COS cells, the recruitment of DP-NTP to cell borders by the chimera required co-expression of plakoglobin. Plakoglobin and DP-NTP co-immunoprecipitated when extracted from L-cells, and yeast two hybrid analysis indicated that DP-NTP binds directly to plakoglobin but not Dsg1. These results identify a role for desmoplakin in organizing the desmosomal cadherin–plakoglobin complex and provide new insights into the hierarchy of protein interactions that occur in the desmosomal plaque.Desmosomes are highly organized adhesive intercellular junctions that couple intermediate filaments to the cell surface at sites of cell–cell adhesion (Farquhar and Palade, 1963; Staehelin, 1974; Schwarz et al., 1990; Garrod, 1993; Collins and Garrod, 1994; Cowin and Burke, 1996; Kowalczyk and Green, 1996). Desmosomes are prominent in tissues that experience mechanical stress, such as heart and epidermis, and the disruption of desmosomes or the intermediate filament system in these organs has devastating effects on tissue integrity (Steinert and Bale, 1993; Coulombe and Fuchs, 1994; Fuchs, 1994; McLean and Lane, 1995; Stanley, 1995; Bierkamp et al., 1996; Ruiz et al., 1996). Desmosomes are highly insoluble structures that can withstand harsh denaturing conditions (Skerrow and Matoltsy, 1974; Gorbsky and Steinberg, 1981; Jones et al., 1988; Schwarz et al., 1990). This property of desmosomes facilitated early identification of desmosomal components but has impaired subsequent biochemical analysis of the protein complexes that form between desmosomal components. Ultrastructurally, desmosomes contain a core region that includes the plasma membranes of adjacent cells and a cytoplasmic plaque that anchors intermediate filaments to the plasma membrane. The plaque can be further divided into an outer dense plaque subjacent to the plasma membrane and an inner dense plaque through which intermediate filaments appear to loop.Molecular genetic analysis has revealed that the desmosomal glycoproteins, the desmogleins and desmocollins, are members of the cadherin family of cell–cell adhesion molecules (for review see Buxton et al., 1993, 1994; Cowin and Mechanic, 1994; Kowalczyk et al., 1996). The classical cadherins, such as E-cadherin, mediate calcium-dependent, homophilic cell–cell adhesion (Nagafuchi et al., 1987). The mechanism by which the desmosomal cadherins mediate cell–cell adhesion remains elusive (Amagai et al., 1994; Chidgey et al., 1996; Kowalczyk et al., 1996), although heterophilic interactions have recently been detected between desmogleins and desmocollins (Chitaev and Troyanovsky, 1997). Both classes of the desmosomal cadherins associate with the cytoplasmic plaque protein plakoglobin (Kowalczyk et al., 1994; Mathur et al., 1994; Roh and Stanley, 1995b ; Troyanovsky et al., 1994), which is part of a growing family of proteins that share a repeated motif first identified in the Drosophila protein Armadillo (Peifer and Wieschaus, 1990). This multigene family also includes the desmosomal proteins band 6/plakophilin 1, plakophilin 2a and 2b, and p0071, which are now considered to comprise a subclass of the armadillo family of proteins (Hatzfeld et al., 1994; Heid et al., 1994; Schmidt et al., 1994; Hatzfeld and Nachtsheim, 1996; Mertens et al., 1996).The most abundant desmosomal plaque protein is desmoplakin, which is predicted to be a homodimer containing two globular end domains joined by a central α-helical coiled-coil rod domain (O''Keefe et al., 1989; Green et al., 1990; Virata et al., 1992). Previous studies have demonstrated that the carboxyl-terminal domain of desmoplakin interacts with intermediate filaments (Stappenbeck and Green, 1992; Stappenbeck et al., 1993; Kouklis et al., 1994; Meng et al., 1997), and the amino-terminal domain of desmoplakin is required for desmoplakin localization to the desmosomal plaque (Stappenbeck et al., 1993). Direct evidence supporting a role for desmoplakin in intermediate filament attachment to desmosomes was provided recently when expression of an amino-terminal polypeptide of desmoplakin was found to displace endogenous desmoplakin from cell borders and disrupt intermediate filament attachment to the cell surface in A431 epithelial cell lines (Bornslaeger et al., 1996).The classical cadherins, such as E-cadherin, bind directly to both β-catenin and plakoglobin (Aberle et al., 1994; Jou et al., 1995; for review see Cowin and Burke, 1996). β-Catenin is also an armadillo family member (McCrea et al., 1991; Peifer et al., 1992), and both plakoglobin and β-catenin bind directly to α-catenin (Aberle et al., 1994, 1996; Jou et al., 1995; Sacco et al., 1995; Obama and Ozawa, 1997). α-Catenin is a vinculin homologue (Nagafuchi et al., 1991) and associates with both α-actinin and actin (Knudson et al., 1995; Rimm et al., 1995; Nieset et al., 1997). Through interactions with β- and α-catenin, E-cadherin is coupled indirectly to the actin cytoskeleton, and this linkage is required for the adhesive activity of E-cadherin (Ozawa et al., 1990; Shimoyama et al., 1992). In addition, E-cadherin association with plakoglobin appears to be required for assembly of desmosomes (Lewis et al., 1997), underscoring the importance of E-cadherin in the overall program of intercellular junction assembly. However, the hierarchy of molecular interactions that couple the desmosomal cadherins to the intermediate filament cytoskeleton is largely unknown, although the desmocollin cytoplasmic domain appears to play an important role in recruiting components of the desmosomal plaque (Troyanovsky et al., 1993, 1994). Since desmosomal cadherins form complexes with plakoglobin and because the amino-terminal domain of desmoplakin is required for desmoplakin localization at desmosomes, we hypothesized that the amino-terminal domain of desmoplakin interacts with the desmosomal cadherin– plakoglobin complex.In previous studies, we used L-cell fibroblasts to characterize plakoglobin interactions with the cytoplasmic domains of the desmosomal cadherins and found that the desmosomal cadherins regulate plakoglobin metabolic stability (Kowalczyk et al., 1994) but do not mediate homophilic adhesion (Kowalczyk et al., 1996). To test the ability of the desmoplakin amino-terminal domain to interact with the desmosomal cadherin–plakoglobin complex, we established a series of L-cell lines expressing the desmosomal cadherins in the presence or absence of a desmoplakin amino-terminal polypeptide (DP-NTP).¹ The results indicate that one important function of the desmoplakin amino-terminal domain is to cluster desmosomal cadherin–plakoglobin complexes. In addition, DP-NTP and plakoglobin were found to form complexes that could be co-immunoprecipitated from L-cell lysates. Using the yeast two hybrid system, DP-NTP was found to bind directly to plakoglobin but not Dsg1. These data suggest that plakoglobin couples the amino-terminal domain of desmoplakin to the desmosomal cadherins and that desmoplakin plays an important role in organizing the desmosomal cadherin–plakoglobin complex into discrete plasma membrane domains.     

The Cell Adhesion Molecule DdCAD-1 in Dictyostelium Is Targeted to the Cell Surface by a Nonclassical Transport Pathway Involving Contractile Vacuoles

DdCAD-1 is a 24-kD Ca²⁺-dependent cell– cell adhesion molecule that is expressed soon after the initiation of development in Dictyostelium cells. DdCAD-1 is present on the cell surface as well as in the cytosol. However, the deduced amino acid sequence of DdCAD-1 lacks a hydrophobic signal peptide or any predicted transmembrane domain, suggesting that it may be presented on the cell surface via a nonclassical transport mechanism. Here we report that DdCAD-1 is transported to the cell surface via contractile vacuoles, which are normally involved in osmoregulation. Immunofluorescence microscopy and subcellular fractionation revealed a preferential association of DdCAD-1 with contractile vacuoles. Proteolytic treatment of isolated contractile vacuoles degraded vacuole-associated calmodulin but not DdCAD-1, demonstrating that DdCAD-1 was present in the lumen. The use of hyperosmotic conditions that suppress contractile vacuole activity led to a dramatic decrease in DdCAD-1 accumulation on the cell surface and the absence of cell cohesiveness. Shifting cells back to a hypotonic condition after hypertonic treatments induced a rapid increase in DdCAD-1–positive contractile vacuoles, followed by the accumulation of DdCAD-1 on the cell membrane. 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole, a specific inhibitor of vacuolar-type H⁺-ATPase and thus of the activity of contractile vacuoles, also inhibited the accumulation of DdCAD-1 on the cell surface. Furthermore, an in vitro reconstitution system was established, and isolated contractile vacuoles were shown to import soluble DdCAD-1 into their lumen in an ATP-stimulated manner. Taken together, these data provide the first evidence for a nonclassical protein transport mechanism that uses contractile vacuoles to target a soluble cytosolic protein to the cell surface.The cellular slime mold Dictyostelium discoideum transits from the solitary amoeboid state to an organized multicellular structure during development. This process is initiated in cells upon the depletion of nutrients, leading to the expression of many developmentally regulated genes and the chemotactic migration of cells in response to extracellular cAMP. Cells stream in concentric rings and/or spirals toward aggregation centers, giving rise to multicellular entities called pseudoplasmodia or slugs. The migrating slugs eventually culminate in the formation of fruiting bodies consisting of primarily spores and stalk cells (for review see Loomis, 1975).Multicellularity during development is maintained by the expression of cell–cell adhesion molecules, which fall into two broad categories based on their sensitivity to EDTA (for reviews see Gerisch, 1980; Siu et al., 1988; Siu, 1990; Fontana, 1995; Bozzaro and Ponte, 1995). There are two types of EDTA-sensitive cell adhesion sites. The EDTA/EGTA-sensitive cell adhesion sites, also known as contact sites B, are mediated by the Ca²⁺-dependent cell adhesion molecule gp24/DdCAD-1 (Knecht et al., 1987; Brar and Siu, 1993), while the EDTA-sensitive/EGTA- resistant sites are probably mediated by a Mg²⁺-dependent cell adhesion molecule (Fontana, 1993). The molecular nature of the latter sites is not yet known. Both types of adhesion sites are responsible for cell–cell interactions in the early stages of development. Coinciding with the aggregation stage is the rapid accumulation of the cell adhesion molecule gp80, which mediates the EDTA-resistant cell adhesion sites or contact sites A (Muller and Gerisch, 1978; Siu et al., 1985; Kamboj et al., 1988, 1989). In postaggregation stages, the EDTA-resistant adhesion sites are mediated by the membrane glycoprotein gp150 (Geltosky et al., 1979; Siu et al., 1983; Gao et al., 1992).DdCAD-1 is expressed by cells soon after the initiation of development (Knecht et al., 1987). Antibodies raised against gel-purified DdCAD-1 specifically inhibit the EDTA/EGTA-sensitive cell–cell adhesion sites and block development (Loomis, 1988). We have purified DdCAD-1 to homogeneity and demonstrated that labeled soluble DdCAD-1 binds to cells in an EDTA/EGTA-sensitive manner (Brar and Siu, 1993). Binding of DdCAD-1 to cells is prevented when cells are precoated with anti– DdCAD-1 antibodies, consistent with a homophilic mode of interaction. In addition, binding of DdCAD-1 to cells inhibits cell reassociation, indicating that it contains only a single cell binding site.Recent cloning of the DdCAD-1 cDNA predicts a protein of 23,924 daltons (Wong et al., 1996). The deduced amino acid sequence of DdCAD-1 shows significant sequence similarities with members of the cadherin family, and it contains a Ca²⁺-binding motif residing in the carboxy-terminal region. Indeed, Ca²⁺ overlay experiments have shown that DdCAD-1 is a Ca²⁺-binding protein with multiple binding sites (Brar and Siu, 1993; Wong et al., 1996). It is therefore conceivable that DdCAD-1 is a primitive member of the cadherin superfamily and it may mediate cell–cell adhesion in a manner similar to that of cadherins (Shapiro et al., 1995; Nagar et al., 1996). Another novel feature of the predicted sequence is that it lacks an amino-terminal hydrophobic signal peptide or a transmembrane domain, suggesting that DdCAD-1 is a soluble protein. Consistent with this observation, both subcellular fractionation and immunofluorescence microscopy have revealed a predominant cytoplasmic localization of DdCAD-1, indicating that 60–80% of DdCAD-1 is soluble (Brar and Siu, 1993; Sesaki and Siu, 1996). However, IgG binding and capping experiments clearly demonstrate that a substantial amount of DdCAD-1 is present on the cell surface (Brar and Siu, 1993; Wong et al., 1996). Interestingly, DdCAD-1 undergoes rapid translocation from the cytoplasm to the plasma membrane in the preaggregation stage of development (Sesaki and Siu, 1996), and then it becomes concentrated on filopodial structures and in cell– cell contact regions. These observations thus raise the question of how DdCAD-1 is transported and anchored to the cell surface.In this report we present morphological and biochemical evidence that DdCAD-1 is transported to the cell surface from the cytosol via contractile vacuoles, which is known so far to function exclusively in osmoregulation in cells. Furthermore, we show that isolated contractile vacuoles selectively take up soluble DdCAD-1 into their lumen in a cell-free system. Our results demonstrate, for the first time, a protein targeting function for contractile vacuoles and a novel nonclassical protein transport mechanism.     

ERS-24, a Mammalian v-SNARE Implicated in Vesicle Traffic between the ER and the Golgi

We report the identification and characterization of ERS-24 (Endoplasmic Reticulum SNARE of 24 kD), a new mammalian v-SNARE implicated in vesicular transport between the ER and the Golgi. ERS24 is incorporated into 20S docking and fusion particles and disassembles from this complex in an ATP-dependent manner. ERS-24 has significant sequence homology to Sec22p, a v-SNARE in Saccharomyces cerevisiae required for transport between the ER and the Golgi. ERS-24 is localized to the ER and to the Golgi, and it is enriched in transport vesicles associated with these organelles.Newly formed transport vesicles have to be selectively targeted to their correct destinations, implying the existence of a set of compartment-specific proteins acting as unique receptor–ligand pairs. Such proteins have now been identified (Söllner et al., 1993a ; Rothman, 1994): one partner efficiently packaged into vesicles, termed a v-SNARE,¹ and the other mainly localized to the target compartment, a t-SNARE. Cognate pairs of v- and t-SNAREs, capable of binding each other specifically, have been identified for the ER–Golgi transport step (Lian and Ferro-Novick, 1993; Søgaard et al., 1994), the Golgi–plasma membrane transport step (Aalto et al., 1993; Protopopov et al., 1993; Brennwald et al., 1994) in Saccharomyces cerevisiae, and regulated exocytosis in neuronal synapses (Söllner et al., 1993a ; for reviews see Scheller, 1995; Südhof, 1995). Additional components, like p115, rab proteins, and sec1 proteins, appear to regulate vesicle docking by controlling the assembly of SNARE complexes (Søgaard et al., 1994; Lian et al., 1994; Sapperstein et al., 1996; Hata et al., 1993; Pevsner et al., 1994).In contrast with vesicle docking, which requires compartment-specific components, the fusion of the two lipid bilayers uses a more general machinery derived, at least in part, from the cytosol (Rothman, 1994), which includes an ATPase, the N-ethylmaleimide–sensitive fusion protein (NSF) (Block et al., 1988; Malhotra et al., 1988), and soluble NSF attachment proteins (SNAPs) (Clary et al., 1990; Clary and Rothman, 1990; Whiteheart et al., 1993). Only the assembled v–t-SNARE complex provides high affinity sites for the consecutive binding of three SNAPs (Söllner et al., 1993b ; Hayashi et al., 1995) and NSF. When NSF is inactivated in vivo, v–t-SNARE complexes accumulate, confirming that NSF is needed for fusion after stable docking (Søgaard et al., 1994).The complex of SNAREs, SNAPs, and NSF can be isolated from detergent extracts of cellular membranes in the presence of ATPγS, or in the presence of ATP but in the absence of Mg²⁺, and sediments at ∼20 Svedberg (20S particle) (Wilson et al., 1992). In the presence of MgATP, the ATPase of NSF disassembles the v–t-SNARE complex and also releases SNAPs. It seems likely that this step somehow initiates fusion.To better understand vesicle flow patterns within cells, it is clearly of interest to identify new SNARE proteins. Presently, the most complete inventory is in yeast, but immunolocalization is difficult in yeast compared with animal cells, and many steps in protein transport have been reconstituted in animal extracts (Rothman, 1992) that have not yet been developed in yeast. Therefore, it is important to create an inventory of SNARE proteins in animal cells. The most unambiguous and direct method for isolating new SNAREs is to exploit their ability to assemble together with SNAPs and NSF into 20S particles and to disassemble into subunits when NSF hydrolyzes ATP. Similar approaches have already been successfully used to isolate new SNAREs implicated in ER to Golgi (Søgaard et al., 1994) and intra-Golgi transport (Nagahama et al., 1996), in addition to the original discovery of SNAREs in the context of neurotransmission (Söllner et al., 1993a ).Using this method, we now report the isolation and detailed characterization of ERS-24 (Endoplasmic Reticulum SNARE of 24 kD), a new mammalian v-SNARE that is localized to the ER and Golgi. ERS-24 is found in transport vesicles associated with the transitional areas of the ER and with the rims of Golgi cisternae, suggesting a role for ERS-24 in vesicular transport between these two compartments.     

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.     

A Novel Class of RanGTP Binding Proteins

The importin-α/β complex and the GTPase Ran mediate nuclear import of proteins with a classical nuclear localization signal. Although Ran has been implicated also in a variety of other processes, such as cell cycle progression, a direct function of Ran has so far only been demonstrated for importin-mediated nuclear import. We have now identified an entire class of ∼20 potential Ran targets that share a sequence motif related to the Ran-binding site of importin-β. We have confirmed specific RanGTP binding for some of them, namely for two novel factors, RanBP7 and RanBP8, for CAS, Pse1p, and Msn5p, and for the cell cycle regulator Cse1p from Saccharomyces cerevisiae. We have studied RanBP7 in more detail. Similar to importin-β, it prevents the activation of Ran''s GTPase by RanGAP1 and inhibits nucleotide exchange on RanGTP. RanBP7 binds directly to nuclear pore complexes where it competes for binding sites with importin-β, transportin, and apparently also with the mediators of mRNA and U snRNA export. Furthermore, we provide evidence for a Ran-dependent transport cycle of RanBP7 and demonstrate that RanBP7 can cross the nuclear envelope rapidly and in both directions. On the basis of these results, we propose that RanBP7 might represent a nuclear transport factor that carries an as yet unknown cargo, which could apply as well for this entire class of related RanGTP-binding proteins.The nuclear pore complexes (NPC)¹ are the sites where the exchange of macromolecules between nucleus and cytoplasm occurs (Feldherr et al., 1984). Transport through the NPCs is bidirectional and comprises a multitude of substrates. Small molecules can passively diffuse through the NPC. The transport of proteins and RNAs >40–60 kD is, however, generally an active process, i.e., it is energy dependent (Newmeyer et al., 1986) and mediated by saturable transport receptors (Goldfarb et al., 1986; Michaud and Goldfarb, 1991; Jarmolowski et al., 1994). To accomplish multiple rounds of transport, these transport receptors are thought to shuttle between nucleus and cytoplasm (Goldfarb et al., 1986). An import receptor, for example, has to bind its import substrate initially in the cytoplasm, release it after NPC passage into the nucleus, and return to the cytoplasm without the cargo. Conversely, an export factor has to bind the export substrate only in the nucleus and on the way out. This model predicts asymmetry in these transport cycles and implies that the binding of the transport receptor to its cargo is regulated by the different environments of nucleus and cytoplasm.The nuclear import of proteins with a classical nuclear localization signal (NLS) is to date the best characterized nucleocytoplasmic transport pathway (for reviews see Görlich and Mattaj, 1996; Koepp and Silver, 1996; Schlenstedt, 1996). The signal contains one or more clusters of basic amino acids (for review see Dingwall and Laskey, 1991) and is recognized by the importin-α/β complex (for references see Sweet and Gerace, 1995; Panté and Aebi, 1996). The α subunit provides the NLS binding site (Imamoto et al., 1995; Weis et al., 1995) and is therefore also called the NLS receptor (Adam and Gerace, 1991). The β subunit accounts for the interaction with the NPC (Görlich et al., 1995; Moroianu et al., 1995) and carries importin-α with the NLS substrate into the nucleus. The translocation into the nucleus is terminated by the disassembly of the importin complex, and both subunits are returned probably separately to the cytoplasm. Importin-α interacts with -β via its importin-β binding domain (IBB domain; Görlich et al., 1996a ; Weis et al., 1996a ). Binding to importin-β with an IBB domain is sufficient for nuclear entry, and the IBB domain can therefore be regarded as the nuclear targeting signal of importin-α. The export domain of importin-α has not yet been identified, but it is distinct from the IBB domain.The small GTPase Ran (Drivas et al., 1990; Bischoff and Ponstingl, 1991b ; Belhumeur et al., 1993) plays a key role in NLS-dependent protein import (Melchior et al., 1993; Moore and Blobel, 1993). GTP hydrolysis by Ran is absolutely essential for import (Melchior et al., 1993; Moore and Blobel, 1993; Schlenstedt et al., 1995a ; Palacios et al., 1996) and is possibly even its sole source of energy (Weis et al., 1996b ). Although the molecular mechanism of import is far from being understood, it appears that Ran fulfils at least two distinct functions in this process: first, Ran''s GTP cycle probably drives translocation into the nucleus (Melchior et al., 1993; Moore and Blobel, 1993; Weis et al., 1996b ), which appears to involve the binding of (cytoplasmic) RanGDP to the NPC, followed by nucleotide exchange and GTP hydrolysis, but it does not involve binding of RanGTP to importin-β (Görlich et al., 1996b ). Unfortunately, nothing is known of how Ran''s GTP cycle would translate into a directed movement through the NPC. Secondly, Ran regulates the interaction between importin-α and -β (Rexach and Blobel, 1995; Chi et al., 1996; Görlich et al., 1996b ). Binding of RanGTP to importin-β disassembles the importin-α/β complex at the nuclear side of the NPC, thereby terminating translocation (Görlich et al., 1996b ). The asymmetric distribution of Ran''s principal GDP/GTP exchange factor (RCC1; Bischoff and Ponstingl, 1991a ) and GTPase activating protein (RanGAP1, or RNA1 in yeast; Bischoff et al., 1995a ; Becker et al., 1995) crucially determines where the importin heterodimer can form and where it is forced to dissociate. RCC1 is a nuclear, chromatin-bound protein (Ohtsubo et al., 1987, 1989) that generates RanGTP in the nucleus, whereas free RanGTP is depleted from the cytoplasm by RanGAP1, which is excluded from the nucleoplasm (Hopper et al., 1990; Matunis et al., 1996; Mahajan et. al, 1997). Thus, low RanGTP levels in the cytoplasm allow importin-α to bind -β, and the high RanGTP concentration in the nuclear compartment dissociates the importin complex. The concentration of free RanGTP can, in this model, be regarded as a marker for cytoplasmic identity (low RanGTP) and nuclear identity (high RanGTP), which is “sensed” by the Ran-binding site in importin-β.It is likely that at least some properties of importin-β are shared by the mediators of the other nucleocytoplasmic transport pathways. This is emphasized by the recent identification of the importin-β–related transportin (Pollard et al., 1996) as an import receptor recognizing the M9 domain, the nuclear targeting signal in hnRNP A1 (Michael et al., 1995), and of yeast transportin (Kap 104p) as an import receptor for mRNA binding proteins (Aitchison et al., 1996). Furthermore, importin-β or its NPC-binding domain cross-compete with other pathways, such as M9-dependent import, NES-mediated nuclear export, and the export of mRNA and U snRNA (Kutay et al., 1997). This would suggest that these other transport receptors share at least some binding sites at the NPC and take a similar path through the nuclear pore complex as importin-β.In addition to importin-β, a number of other Ran-binding proteins are detectable in eukaryotic cells, e.g., in overlay blots using Ran γ-[³²P]GTP as a probe. These can be grouped into two classes (Lounsbury et al., 1994, 1996): first, those with a RanBP1 homology domain including the Ran binding protein 1 (RanBP1) itself (Coutavas et al., 1993; Bischoff et al., 1995b ) and the nuclear pore protein RanBP2, which has four RanBP1 homology domains (Wu et al., 1995; Yokoyama et al., 1995). Their binding to Ran can be competed by RanBP1. Second, importin-β and so far unidentified protein(s) of ∼120 kD whose Ran-binding is competed by importin-β but not by excess of RanBP1 (Lounsbury et al., 1994, 1996). Both RanBP1 and importin-β inhibit the nucleotide exchange on RanGTP (Coutavas et al., 1993; Lounsbury et al., 1994, 1996; Bischoff et al., 1995b ; Görlich et al., 1996b ). However, they do not cross-compete with each other for Ran binding but instead bind to different, nonoverlapping sites on Ran (Chi et al., 1996; Kutay et al., 1997; Lounsbury and Macara, 1997). Another striking difference is that RanBP1 facilitates the activation of Ran''s GTPase by RanGAP1 (Beddow et al., 1995; Bischoff et al., 1995b ), whereas the importin-β/RanGTP complex is entirely GAP resistant (Floer and Blobel, 1996; Görlich et al., 1996b ).Although a direct involvement of Ran has so far only been demonstrated in the importin-dependent transport pathway, perturbations in the Ran system have severe effects on a great variety of cellular functions, such as RNA processing, RNA export, regulation of chromosome structure, cell cycle progression, initiation of replication, microtubule structure, etc. (for review see Dasso, 1993; Sazer, 1996). One could argue that these effects are all secondary consequences from an impaired NLS-dependent protein import. However, it is also possible that these defects are more direct and that eukaryotic cells contain many immediate targets of Ran function.Here we describe a novel superfamily of Ran-binding proteins, which includes about a dozen factors in yeast and probably even more in higher eukaryotes. The members of this superfamily share with importin-β an NH₂-terminal sequence motif that appears to account for RanGTP binding. Indeed we could confirm the interaction with Ran for the following factors: RanBP7 and RanBP8, two novel, related proteins described here, Cse1p, a cell cycle regulator in yeast, CAS, which is required for apoptosis in cultured human cells, and for Msn5p and Pse1p from yeast. Of these we have characterized RanBP7 and RanBP8 in more detail. Both resemble closely importin-β in their interaction with Ran, and both bind directly to nuclear pore complexes. RanBP7 can cross the nuclear membrane rapidly and in both directions. We provide evidence for a transport cycle in which RanBP7 first enters the nucleus, binds RanGTP inside the nucleus as a prerequisite for rapid re-export to the cytoplasm, after which the RanBP7/RanGTP complex becomes finally disassembled by the concerted action of RanBP1 and RanGAP1 in the cytoplasm. We propose that during these transport cycles, RanBP7 would normally carry an as yet unidentified cargo. This means, RanBP7 and possibly also the other members of the RanBP7/Cse1p/ importin-β superfamily could function as transport receptors that shuttle between nucleus and cytoplasm. RanBP7 and importin-β form an abundant, heterodimeric complex in the cytoplasm that appears to have a function different from nuclear import of proteins with a classical NLS. It might be a way to regulate either RanBP7 or importin-β function. Alternatively, the RanBP7/importin-β complex might constitute an import receptor with a substrate specificity different from that of the importin-α/β complex.     

Sec35p,a Novel Peripheral Membrane Protein,Is Required for ER to Golgi Vesicle Docking

SEC35 was identified in a novel screen for temperature-sensitive mutants in the secretory pathway of the yeast Saccharomyces cerevisiae (Wuestehube et al., 1996. Genetics. 142:393–406). At the restrictive temperature, the sec35-1 strain exhibits a transport block between the ER and the Golgi apparatus and accumulates numerous vesicles. SEC35 encodes a novel cytosolic protein of 32 kD, peripherally associated with membranes. The temperature-sensitive phenotype of sec35-1 is efficiently suppressed by YPT1, which encodes the rab-like GTPase required early in the secretory pathway, or by SLY1-20, which encodes a dominant form of the ER to Golgi target -SNARE–associated protein Sly1p. Weaker suppression is evident upon overexpression of genes encoding the vesicle-SNAREs SEC22, BET1, or YKT6. The cold-sensitive lethality that results from deleting SEC35 is suppressed by YPT1 or SLY1-20. These genetic relationships suggest that Sec35p acts upstream of, or in conjunction with, Ypt1p and Sly1p as was previously found for Uso1p. Using a cell-free assay that measures distinct steps in vesicle transport from the ER to the Golgi, we find Sec35p is required for a vesicle docking stage catalyzed by Uso1p. These genetic and biochemical results suggest Sec35p acts with Uso1p to dock ER-derived vesicles to the Golgi complex.Protein transport through the secretory pathway occurs via transport vesicles under the direction of a large set of protein components (Rothman, 1994). The process can be divided into three stages: (a) vesicle budding, (b) vesicle docking, and (c) membrane fusion, with distinct sets of proteins mediating each phase. The budding step involves recruitment of coat proteins to the membrane and culminates with the release of coated vesicles (Schekman and Orci, 1996). The docking reaction is likely to require a set of integral membrane proteins on the vesicle and target membranes, termed v-SNAREs¹ and t-SNAREs (vesicle- and target membrane-soluble N-ethylmaleimide–sensitive fusion protein [NSF] attachment protein [SNAP] receptors, respectively), that are thought to confer specificity through their pair-wise interactions (Söllner et al., 1993b ). Small GTP-binding proteins of the rab family also assist in the docking process (Ferro-Novick and Novick, 1993), but their precise function is not known. The fusion step ensues after docking and results in the delivery of the vesicular cargo to the next compartment in the secretory pathway.Vesicular transport from the ER to the Golgi apparatus in the yeast Saccharomyces cerevisiae has been extensively characterized. Transport vesicle budding involves the assembly of the COPII coat, composed of the Sec13p/Sec31p (Pryer et al., 1993; Salama et al., 1993; Barlowe et al., 1994) and Sec23p/Sec24p heterodimers (Hicke and Schekman, 1989; Hicke et al., 1992), under the direction of an integral membrane protein, Sec12p (Nakano et al., 1988; Barlowe and Schekman, 1993), a small GTPase, Sar1p (Nakano and Muramatsu, 1989), and a multidomain protein, Sec16p (Espenshade et al., 1995; Shaywitz et al., 1997). Docking is thought to require a tethering event mediated by Uso1p (Cao et al., 1998), the yeast homologue of mammalian p115 (Barroso et al., 1995; Sapperstein et al., 1995), followed by or concurrent with the interaction of a set of ER to Golgi v-SNAREs, Bet1p, Bos1p, Sec22p (Newman and Ferro-Novick, 1987; Newman et al., 1990; Ossig et al., 1991; Shim et al., 1991; Søgaard et al., 1994) and perhaps Ykt6p (Søgaard et al., 1994; McNew et al., 1997), with the cognate t-SNARE on the Golgi, Sed5p (Hardwick and Pelham, 1992). For some time it was thought that fusion may be initiated by disassembly of the v/t-SNARE complex (Söllner et al., 1993a ) by yeast SNAP, Sec17p, (Griff et al., 1992) and NSF, Sec18p (Eakle et al., 1988; Wilson et al., 1989). However, this concept has been challenged by studies with a yeast system that reconstitutes homotypic vacuolar fusion, which suggests the action of Sec18p is before vesicle docking (Mayer et al., 1996; Mayer and Wickner, 1997). In addition, a prefusion role for NSF has been supported by the recent finding that liposomes bearing SNAREs alone can fuse in the absence of NSF (Weber et al., 1998).Several proteins involved in the regulation of yeast ER to Golgi v/t-SNARE complex assembly have been identified, including Ypt1p, Uso1p, and Sly1p. Ypt1p is a member of the rab family of small GTP-binding proteins that have been identified as important components of almost every stage in the secretory pathway (Ferro-Novick and Novick, 1993). Hydrolysis of GTP by rab-like proteins has been hypothesized to provide the regulatory switch that controls the fidelity of vesicular transport (Bourne, 1988). A second protein, Uso1p (Nakajima et al., 1991), appears to function in the same pathway as Ypt1p (Sapperstein et al., 1996), and both proteins have been demonstrated to be essential for SNARE complex assembly (Søgaard et al., 1994; Sapperstein et al., 1996; Lupashin and Waters, 1997). The third protein, Sly1p, is associated with the t-SNARE Sed5p (Søgaard et al., 1994). SLY1 is an essential gene in yeast (Dascher et al., 1991; Ossig et al., 1991), and Sly1p is required for ER to Golgi transport in vitro (Lupashin et al., 1996) and in vivo (Ossig et al., 1991). However, several lines of evidence, particularly from Sly1p homologues in other organisms, indicate that Sly1p may also function as a negative regulator of v/t-SNARE complex assembly, perhaps by preventing the association of the v- and t-SNAREs (Hosono et al., 1992; Pevsner et al., 1994; Schulze et al., 1994). A dominant allele of SLY1, termed SLY1-20, is capable of suppressing mutations in YPT1 and USO1, including complete deletions (Dascher et al., 1991; Sapperstein et al., 1996). Thus, in the presence of Sly1-20p, two components required for SNARE complex assembly are no longer essential. We have proposed a model (Sapperstein et al., 1996; Lupashin and Waters, 1997) in which Ypt1p and Uso1p function to relieve the inhibitory action of Sly1p on SNARE complex assembly. In this model Sly1-20p can be thought of as a noninhibitory form of SLY1 that renders Ypt1p and Uso1p superfluous.We believe that the ability of SLY1-20 to suppress defects in upstream docking regulators can be used to identify additional components involved in the regulation of vesicular docking. We have undertaken a genetic screen (to be presented elsewhere) to isolate novel components in this pathway which, when mutated, depend upon Sly1-20p for viability. In the course of this work, we discovered that two recently identified mutants, sec34 and sec35, can be suppressed by SLY1-20 and thus satisfy the criterion of our screen. These mutants were isolated in a novel screen to identify components involved in transport at any step between the ER and the trans-Golgi network (i.e., the Kex2p compartment) in yeast (Wuestehube et al., 1996). Both sec34 and sec35 accumulate the core-glycosylated form of secretory proteins at the nonpermissive temperature, indicating a block in ER to Golgi transport. Furthermore, electron microscopy indicated that both sec34 and sec35 accumulate numerous vesicles upon shift to the restrictive temperature (Wuestehube et al., 1996), a hallmark of genes whose protein products are involved in the docking or fusion phase of transport (Kaiser and Schekman, 1990). In this report we describe the cloning of SEC35 and analysis of its genetic interactions with other secretory genes. Strong genetic interaction between SEC35 and SLY1, YPT1, and USO1 suggests that Sec35p may function in vesicle docking. To test this possibility, we devised an in vitro transport assay that depends on the addition of purified Sec35p and Uso1p. Vesicles synthesized in the absence of functional Sec35p do not fuse with the Golgi compartment and remain as freely diffusible intermediates. Upon addition of Sec35p and Uso1p, vesicles dock to the Golgi and proceed to membrane fusion. Requirements for Sec35p at the vesicle docking step correlates our genetic experiments with the biochemically distinguishable steps of vesicle docking and membrane fusion.     

Three-dimensional Structure of Acanthamoeba castellanii Myosin-IB (MIB) Determined by Cryoelectron Microscopy of Decorated Actin Filaments

The Acanthamoeba castellanii myosin-Is were the first unconventional myosins to be discovered, and the myosin-I class has since been found to be one of the more diverse and abundant classes of the myosin superfamily. We used two-dimensional (2D) crystallization on phospholipid monolayers and negative stain electron microscopy to calculate a projection map of a “classical” myosin-I, Acanthamoeba myosin-IB (MIB), at ∼18 Å resolution. Interpretation of the projection map suggests that the MIB molecules sit upright on the membrane. We also used cryoelectron microscopy and helical image analysis to determine the three-dimensional structure of actin filaments decorated with unphosphorylated (inactive) MIB. The catalytic domain is similar to that of other myosins, whereas the large carboxy-terminal tail domain differs greatly from brush border myosin-I (BBM-I), another member of the myosin-I class. These differences may be relevant to the distinct cellular functions of these two types of myosin-I. The catalytic domain of MIB also attaches to F-actin at a significantly different angle, ∼10°, than BBM-I. Finally, there is evidence that the tails of adjacent MIB molecules interact in both the 2D crystal and in the decorated actin filaments.The myosin superfamily consists of at least 12 distinct classes that vary both in the sequence of their conserved myosin catalytic domains as well as in their unique tails (Mooseker and Cheney, 1995; Sellers and Goodson, 1995). For many years the only known myosins were the double-headed, filament-forming myosins found in muscle (conventional myosins or myosins-II). The remaining classes of myosin have been termed “unconventional myosins” to differentiate them from the myosins-II. Probably the most thoroughly studied class of unconventional myosins is the myosin-I class. These small, single-headed myosins bind to membrane lipids through a basic domain in their tail (for review see Pollard et al., 1991; Mooseker and Cheney, 1995). The first unconventional myosin (and first myosin-I) was isolated from Acanthamoeba castellanii (Pollard and Korn, 1973 a,b), and was purified on the basis of its K⁺, EDTA, and actin-activated MgATPase activities. However, this myosin was unusual in that it had a single heavy chain of ∼140 kD, in contrast to the two ∼200-kD heavy chains of myosin-II (Pollard and Korn, 1973 a).Three isoforms of the classical Acanthamoeba myosins-I are now known: myosins-IA, -IB, and -IC (Maruta and Korn, 1977a ,b; Maruta et al., 1979). Each of the isoforms consists of a conserved myosin catalytic domain, a binding site for one or two light chains, a basic domain, a GPA¹(Q) domain (rich in glycine, proline and alanine [glutamine]), and an scr-homology domain-3 (SH3) domain (Pollard et al., 1991; Mooseker and Cheney, 1995). These myosins-I can associate with membranes or with actin filaments through their tail domains. An electrostatic association of myosin-I with anionic phospholipids and with base-stripped membranes has been shown to occur (Adams and Pollard, 1989; Miyata et al., 1989; Hayden et al., 1990), and this interaction has been mapped to the basic domain (Doberstein and Pollard, 1992). Interestingly, these myosins also contain a second, ATP-insensitive actin binding site (Lynch et al., 1986) enabling them to mediate actin–actin movements (Albanesi et al., 1985; Fujisaki et al., 1985). In myosin-IA (Lynch et al., 1986) and myosin-IC (Doberstein and Pollard, 1992), this binding site was localized to the GPA domain. Acanthamoeba myosins-I have maximal steady-state actin-activated ATPase rates of ∼10–20 s⁻¹ (Pollard and Korn, 1973 b; Albanesi et al., 1983), and an unusual triphasic dependence upon actin concentration (Pollard and Korn, 1973 b; Albanesi et al., 1983). This triphasic activation is due to the actin cross-linking ability imparted by the ATP-insensitive actin binding site on the tail (Albanesi et al., 1985). Analysis of the individual steps in the ATPase cycle by transient kinetics revealed that the mechanism of myosin-IA is similar to slow skeletal muscle myosin, whereas myosin-IB (MIB) is similar to fast skeletal muscle myosin (Ostap and Pollard, 1996). The in vitro motility of myosin-I has also been well characterized (Zot et al., 1992). The maximal rate of filament sliding is ∼0.2 μm s⁻¹. Interestingly, this rate is ∼10–50× slower than the rates observed for skeletal muscle myosin, even though the ATPase rates are comparable.MIB consists of a 125-kD heavy chain and a single 27-kD light chain (Maruta et al., 1979; Jung et al., 1987). This isoform is primarily associated with the plasma membrane as well as vacuolar membranes (Baines et al., 1992). MIB appears to be associated with the plasma membrane at sites of phagocytosis and was concentrated at the tips of pseudopodia (Baines et al., 1992). This localization suggests that MIB may be involved in membrane dynamics at the cell surface. MIB is regulated by heavy chain phosphorylation of serine 411 (Brzeska et al., 1989, 1990), which is located at the actin-binding site (Rayment et al., 1993). Similar to the myosin-I isoforms in Acanthamoeba, heavy chain phosphorylation results in >20-fold activation of the actin-activated myosin-I ATPase activity (Albanesi et al., 1983). This activation is not the result of changes in the binding of myosin-I to F-actin (Albanesi et al., 1983; Ostap and Pollard, 1996). The transient kinetic studies of Ostap and Pollard (1996) suggest that phosphorylation regulates the rate-limiting phosphate release step, the transition from weakly bound intermediates in rapid equilibrium with actin to strongly bound states, capable of sustaining force.Despite the extensive analysis of ameboid myosin-I biochemical properties and in vivo function, there is little structural information on these myosins. The only detailed structural information available for the myosins-I comes from recent electron microscopy studies on brush border myosin-I (BBM-I) (Jontes et al., 1995; Jontes and Milligan, 1997a ,b; Whittaker and Milligan, 1997), a structurally distinct myosin-I subtype. Therefore, we investigated the structure of a “classical,” ameboid-type myosin, Acanthamoeba MIB using electron microscopy. First, electron micrographs of negatively stained two-dimensional (2D) crystals were used to generate a projection map of MIB at ∼18 Å resolution. In addition, we used cryoelectron microscopy and helical image analysis to produce a moderate resolution three-dimensional (3D) map (30 Å) of actin filaments decorated with MIB. These studies enabled us to compare the structure of MIB with BBM-I. The comparison of MIB with BBM-I reveals marked structural differences in the tail domains of these two proteins; MIB appears to have a much shorter “lever arm” and a more compact tail, whereas most of the BBM-I mass is composed of an extended light chain–binding domain (LCBD). In addition, the MIB catalytic domain appears to be slightly tilted compared to BBM-I, with respect to the F-actin axis. Our structural results suggest that these two types of myosin-I may have distinct intracellular functions.     

C-Nap1, a Novel Centrosomal Coiled-Coil Protein and Candidate Substrate of the Cell Cycle–regulated Protein Kinase Nek2

Nek2 (for NIMA-related kinase 2) is a mammalian cell cycle–regulated kinase structurally related to the mitotic regulator NIMA of Aspergillus nidulans. In human cells, Nek2 associates with centrosomes, and overexpression of active Nek2 has drastic consequences for centrosome structure. Here, we describe the molecular characterization of a novel human centrosomal protein, C-Nap1 (for centrosomal Nek2-associated protein 1), first identified as a Nek2-interacting protein in a yeast two-hybrid screen. Antibodies raised against recombinant C-Nap1 produced strong labeling of centrosomes by immunofluorescence, and immunoelectron microscopy revealed that C-Nap1 is associated specifically with the proximal ends of both mother and daughter centrioles. On Western blots, anti–C-Nap1 antibodies recognized a large protein (>250 kD) that was highly enriched in centrosome preparations. Sequencing of overlapping cDNAs showed that C-Nap1 has a calculated molecular mass of 281 kD and comprises extended domains of predicted coiled-coil structure. Whereas C-Nap1 was concentrated at centrosomes in all interphase cells, immunoreactivity at mitotic spindle poles was strongly diminished. Finally, the COOH-terminal domain of C-Nap1 could readily be phosphorylated by Nek2 in vitro, as well as after coexpression of the two proteins in vivo. Based on these findings, we propose a model implicating both Nek2 and C-Nap1 in the regulation of centriole–centriole cohesion during the cell cycle.The serine/threonine kinase NIMA of Aspergillus nidulans is considered the founding member of a family of protein kinases with a possible role in cell cycle regulation (for reviews see Fry and Nigg, 1995; Lu and Hunter, 1995a ; Osmani and Ye, 1996). In A. nidulans, NIMA clearly cooperates with the Cdc2 protein kinase to promote progression into mitosis (Osmani et al., 1991), and overexpression of NIMA in a variety of heterologous species promotes a premature onset of chromosome condensation (O''Connell et al., 1994; Lu and Hunter, 1995b ). This has been interpreted to suggest evolutionary conservation of a pathway involving NIMA-related kinases (for review see Lu and Hunter, 1995a ). Indeed, kinases structurally related to NIMA are present in many species (Fry and Nigg, 1997). However, the only bona fide functional homologue of NIMA so far isolated stems from another filamentous fungus, Neurospora crassa (Pu et al., 1995), and the functional relationship between vertebrate NIMA-related kinases and fungal NIMA remains uncertain.The closest known mammalian relative to NIMA is a kinase termed Nek2 (for NIMA-related kinase 2)¹ (Fry and Nigg, 1997). This kinase undergoes cell cycle–dependent changes in abundance and activity, reminiscent of NIMA (Schultz et al., 1994; Fry et al., 1995). It is highly expressed in male germ cells (Rhee and Wolgemuth, 1997; Tanaka et al., 1997), and data have been reported consistent with a role for Nek2 in meiotic chromosome condensation (Rhee and Wolgemuth, 1997). However, overexpression of active Nek2 in somatic cells has no obvious effect on chromosome condensation; instead, it induces striking alterations in the structure of the centrosome, the principal microtubule-organizing center of mammalian cells (Fry et al., 1998). Furthermore, immunofluorescence microscopy and subcellular fractionation concur to demonstrate that endogenous Nek2 associates with centrosomes, strongly suggesting that one physiological function of this kinase may relate to the centrosome cycle (Fry et al., 1998).The mammalian centrosome is an organelle of about 1 μm in diameter. It comprises two barrel-shaped centrioles that are made of nine short triplet microtubules and are surrounded by an amorphous matrix known as the pericentriolar material (PCM) (for review see Brinkley, 1985; Vorobjev and Nadehzdina, 1987; Kimble and Kuriyama, 1992; Kalt and Schliwa, 1993; Kellogg et al., 1994; Lange and Gull, 1996). Major progress has recently been made with the demonstration that microtubules are nucleated from γ-tubulin–containing ring complexes (γ-TuRCs), which are concentrated within the PCM (Moritz et al., 1995; Zheng et al., 1995). γ-Tubulin forms complexes with Spc97/98, two evolutionarily conserved proteins first identified in budding yeast spindle pole bodies (Geissler et al., 1996; Knop et al., 1997; Stearns and Winey, 1997), and there is also evidence for an important role of pericentrin and other coiled-coil proteins in organizing γ-TuRCs into higher order lattice structures (Doxsey et al., 1994; Dictenberg et al., 1998). However, in spite of this recent progress, it is clear that the inventory of centrosome components is far from complete.Centrosome structure and function is regulated in a cell cycle–dependent manner (for reviews see Mazia, 1987; Kellogg et al., 1994; Tournier and Bornens, 1994). Once in every cell cycle, and beginning around the G1/S transition, centrioles are duplicated (e.g., Kuriyama and Borisy, 1981a ; Vorobjev and Chentsov, 1982; Kochanski and Borisy, 1990; Chrétien et al., 1997). Late in G2, centrosomes then grow in size (a process referred to as maturation) through the recruitment of additional PCM proteins (Rieder and Borisy, 1982; Kalt and Schliwa, 1993; Lange and Gull, 1995). At the G2/M transition, the duplicated centrosomes separate and migrate to opposite ends of the nucleus. Concomitantly, their microtubule-nucleating activities increase dramatically in preparation for spindle formation (McGill and Brinkley, 1975; Snyder and McIntosh, 1975; Gould and Borisy, 1977; Kuriyama and Borisy, 1981b ; for reviews see Brinkley, 1985; Vorobjev and Nadehzdina, 1987; Karsenti, 1991). By what mechanisms these events are controlled remains largely unknown, but data obtained using phosphoepitope-specific antibodies strongly suggest that phosphorylation of centrosomal proteins plays a major role (Vandré et al., 1984, 1986; Centonze and Borisy, 1990). More direct support for this view stems from the observation that cyclin-dependent kinases (CDKs) enhance the microtubule-nucleation activity of centrosomes at the G2/M transition (Verde et al., 1990, 1992; Buendia et al., 1992) and are involved in promoting centrosome separation (Blangy et al., 1995; Sawin and Mitchison, 1995). Similarly, polo-like kinase 1, a cell cycle regulatory kinase structurally distinct from CDKs, has recently been implicated in centrosome maturation (Lane and Nigg, 1996).The precise role of Nek2 at the centrosome remains to be determined, but it is intriguing that overexpression of this kinase in human cells causes a pronounced splitting of centrosomes. This led us to propose that Nek2-dependent phosphorylation of previously unidentified proteins may cause a loss of centriole–centriole cohesion, and that this event might represent an early step in centrosome separation at the G2/M transition (Fry et al., 1998). With the aim of identifying potential substrates (or regulators) of Nek2, we have now performed a yeast two-hybrid screen, using full-length Nek2 as a bait. We report here the molecular characterization of a novel coiled-coil protein that we call C-Nap1 (for centrosomal Nek2-associated protein 1). C-Nap1 represents a core component of the mammalian centrosome and the first candidate substrate for a member of the NIMA protein kinase family to be identified.