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.     

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.     

Brain Myosin V Is a Synaptic Vesicle-associated Motor Protein: Evidence for a Ca2+-dependent Interaction with the Synaptobrevin–Synaptophysin Complex

Brain myosin V is a member of a widely distributed class of unconventional myosins that may be of central importance to organelle trafficking in all eukaryotic cells. Molecular constituents that target this molecular motor to organelles have not been previously identified. Using a combination of immunopurification, extraction, cross-linking, and coprecipitation assays, we demonstrate that the tail domain of brain myosin V forms a stable complex with the synaptic vesicle membrane proteins, synaptobrevin II and synaptophysin. While myosin V was principally bound to synaptic vesicles during rest, this putative transport complex was promptly disassembled upon the depolarization-induced entry of Ca²⁺ into intact nerve endings. Coimmunoprecipitation assays further indicate that Ca²⁺ disrupts the in vitro binding of synaptobrevin II to synaptophysin in the presence but not in the absence of Mg²⁺. We conclude that hydrophilic forces reversibly couple the myosin V tail to a biochemically defined class of organelles in brain nerve terminals.Synaptic vesicles are neuronal organelles that sequester, store, and release neurotransmitters. Functionally mature synaptic vesicles are assembled locally, in the nerve terminal, from at least two different types of precursor vesicles (Okada et al., 1995). Once the mature synaptic vesicle becomes docked at the presynaptic membrane, it is prepared (“primed”) to rapidly discharge its shipment of transmitter into the synaptic cleft upon the arrival of an action potential. This secretory response is triggered by the opening of voltage-regulated Ca²⁺ channels and is the principle mechanism used by neurons to exchange information.A majority of the synaptic vesicles in a nerve terminal are sequestered into clusters that are not immediately available for exocytosis (Landis et al., 1988; Hirokawa et al., 1989). Thus, the initial burst of Ca²⁺-triggered exocytosis is presently understood to result from the fusion of predocked vesicles that had matured to a fusion-competent state before the action potential arrived (Südhof, 1995; Rosenmund and Stevens, 1996). The current working model further interprets the existence of vesicle clusters as a potential reserve pool of synaptic vesicles that may be mobilized in a use-dependent manner to replenish the pool of readily releasable vesicles. Several independent lines of experimental evidence support this scheme. Electrophysiological measurements of membrane fusion report that hippocampal synapses recover from complete synaptic fatigue with a time constant of ∼10 s (Stevens and Tsujimoto, 1995; Rosenmund and Stevens, 1996), suggesting that mechanisms exist for actively replenishing the readily releasable pool of synaptic vesicles. The kinetics of synaptic vesicle recycling indicate that recently retrieved vesicle membranes do not completely fulfill the demand for releasable vesicles during periods of intense secretory activity (Ryan et al., 1993; Ryan and Smith, 1995). This analysis implies that a supply of fresh synaptic vesicles must somehow move from a reserve pool to the presynaptic membrane. In fact, movements of synaptic vesicles have now been observed in a variety of different synaptic preparations (Llinas et al., 1989; Koenig et al., 1993; Henkel et al., 1996). These movements are inconsistent with simple diffusion, since they are reported to be both ATP-dependent and vectorial in nature. Integral synaptic vesicle proteins are transported to the terminal by a microtubule-based superfamily of motor proteins, the kinesins (Okada et al., 1995). However, microtubules do not extend into the cortical cytoskeletal matrix of terminals (Landis et al., 1988; Hirokawa et al., 1989), and the kinesins are rapidly degraded upon their arrival in the terminal (Okada et al., 1995). Moreover, recent imaging studies have shown that intracellular particles move along actin bundles in nerve growth cones, rather than microtubules (Evans and Bridgman, 1995). For these reasons, it seems unlikely that the kinesins move synaptic vesicles between their putative storage sites (the vesicle clusters) and the sites of their release (the active zones). Despite intensive study of the molecular events that govern synaptic vesicle recycling, relatively little is known about the mechanisms that control synaptic vesicle movements within the nerve terminal.Brain myosins V (p190, dilute, myr 6) are presently the leading candidates for a synaptic vesicle motor protein (Langford, 1995). In the nervous system, the mouse dilute (Mercer et al., 1991) and chicken myosin V (p190; Espreafico et al., 1992) proteins are localized within neurons and are more abundant than myr 6, which is concentrated in the dentate gyrus and choroid plexus (Zhao et al., 1996). The myosin superfamily includes at least 11 different molecular motors that possess a conserved motor domain attached to a variety of neck and tail domains with distinguishing structural and biochemical characteristics (Cheney et al., 1993b ; Mooseker and Cheney, 1995). Both the conventional myosins II and unconventional myosins V have been localized within presynaptic terminals (Espreafico et al., 1992; Mochida et al., 1994; see also Miller et al., 1992); form a two-headed structure that is capable of generating mechanochemical force and moving actin filaments; and possess actin-activated Mg²⁺-ATPase activity. In addition, both forms of myosins have neck domains that contain variable numbers of putative regulatory calmodulin light chain binding cassettes, and a flexible coiled-coil stalk that is followed by a tail domain which ends as a large (∼400 amino acids) COOH-terminal globular structure (Espreafico et al., 1992; Cheney et al., 1993a ). Diversification of the intracellular localization and functions of the myosins seems to depend on the COOH-terminal tail which is variable in sequence and length (Cheney et al., 1993b ). Myosins II and V appear to regulate complementary presynaptic functions: actin dynamics and the transport of membrane-bound organelles, respectively. The heavy chains of myosins IIA and IIB bind to acidic phospholipids and self-assemble into contractile filaments on the subplasmallemal surface (Murakami et al., 1995; Verkhovsky et al., 1995), which may play an important role in growth cone motility (Miller et al., 1992), the cycling of cortical actin assembly (Bernstein and Bamburg, 1989), and the regulation of neurotransmitter release (Mochida et al., 1994). In contrast, myosins V are not filament-forming, and they colocalize with intracellular membranes (Cheney et al., 1993a ). The class V myosins may have a role in regulating the extension of growth cone filapodia during neuronal development (Wang et al., 1996), as well as the movement of axoplasmic organelles (Kuznetsov et al., 1992; Bearer et al., 1993; Langford, 1995). A potential role for brain myosin V in organelle trafficking and transport is indirectly supported by the finding that this motor protein is capable of moving relatively large (800 nm) artificial beads in motility assays (Wolenski et al., 1995). Moreover, functional null mutations have established that the class V myosins are required in yeast for the intracellular transport of post-Golgi secretory vesicles (Johnston et al., 1991; Lillie and Brown, 1992; Govindan et al., 1995).Myosin motors bound directly to synaptic vesicles may have an important role in regulating the availability of synaptic vesicles for exocytosis. The present study demonstrates that 2,3-butanedione-2-monoxime (BDM),¹ a pharmacological inhibitor of endogenous myosin Mg²⁺-ATPase activity, significantly decreases the Ca²⁺-dependent release of glutamate from isolated nerve endings (synaptosomes). In addition, our data reveal that a major portion of the myosin V in nerve terminals was bound to synaptic vesicles and that this interaction was regulated by Ca²⁺ concentrations that are of physiological relevance. Cross-linking experiments indicated that myosin V specifically complexes with the synaptic vesicle proteins synaptobrevin and synaptophysin. Interactions between these two integral membrane proteins may impose an important additional constraint on the regulated pathway of secretion (Calakos and Scheller, 1994; Edelmann et al., 1995). We now present evidence that the cytoplasmic domain of the synaptobrevin–synaptophysin complex may also function as a binding partner for myosin V, forming a multimeric complex that shall be referred to as the “myosin V transport complex.”     

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.     

Dynamic Properties of an Inositol 1,4,5-Trisphosphate– and Thapsigargin-insensitive Calcium Pool in Mammalian Cell Lines

The functional characteristics of a nonacidic, inositol 1,4,5-trisphosphate– and thapsigargin-insensitive Ca²⁺ pool have been characterized in mammalian cells derived from the rat pituitary gland (GH3, GC, and GH3B6), the adrenal tissue (PC12), and mast cells (RBL-1). This Ca²⁺ pool is released into the cytoplasm by the Ca²⁺ ionophores ionomycin or A23187 after the discharge of the inositol 1,4,5-trisphosphate–sensitive store with an agonist coupled to phospholipase C activation and/or thapsigargin. The amount of Ca²⁺ trapped within this pool increased significantly after a prolonged elevation of intracellular Ca²⁺ concentration elicited by activation of Ca²⁺ influx. This pool was affected neither by caffeine-ryanodine nor by mitochondrial uncouplers. Probing mitochondrial Ca²⁺ with recombinant aequorin confirmed that this pool did not coincide with mitochondria, whereas its homogeneous distribution across the cytosol, as revealed by confocal microscopy, and its insensitivity to brefeldin A make localization within the Golgi complex unlikely. A proton gradient as the driving mechanism for Ca²⁺ uptake was excluded since ionomycin is inefficient in releasing Ca²⁺ from acidic pools and Ca²⁺ accumulation/release in/from this store was unaffected by monensin or NH₄Cl, drugs known to collapse organelle acidic pH gradients. Ca²⁺ sequestration inside this pool, thus, may occur through a low-affinity, high-capacity Ca²⁺–ATPase system, which is, however, distinct from classical endosarcoplasmic reticulum Ca²⁺–ATPases. The cytological nature and functional role of this Ca²⁺ storage compartment are discussed.The cytosolic free Ca²⁺ concentration ([Ca²⁺]_i)¹ of eukaryotic cells rests in the range of 50–200 nM, i.e., at a very low level, if compared to the Ca²⁺ concentration of physiological media (2 mM). However, the total cellular Ca²⁺ content is closer to this latter value (1–3 mmol/l of cell water). In other words, eukaryotic cells sequester large amounts of Ca²⁺ mainly by uptake inside intracellular Ca²⁺ stores (∼90%) (for reviews see Pozzan et al., 1994; Clapham, 1995).The complexity of intracellular Ca²⁺ stores has been intensively investigated in recent years (for reviews see Meldolesi et al., 1990; Pozzan et al., 1994; Simpson et al., 1995). Attention has been focused mainly on Ca²⁺ stores that are highly dynamic because of their ability to rapidly take up and release Ca²⁺. Ca²⁺ sequestration into these pools depends on Ca²⁺–ATPases, known as sarco/endoplasmic reticulum Ca²⁺–ATPases (SERCAs) (Burk et al., 1989; Bobe et al., 1994; Wuytack et al., 1994). All the SERCA isoforms share the property of being selectively inhibited by thapsigargin (Tg), a tumor-promoting sesquiterpene lactone (Lytton et al., 1991). Tg acts with both high affinity, at nanomolar concentrations, and high specificity, with virtually no effect on the Ca²⁺– or Na⁺/K⁺– ATPase of the plasmalemma.Other drugs, such as 2,5-di(tert-butyl)-1,4-benzohydroquinone (tBHQ) and cyclopiazonic acid (CA), also block SERCAs, albeit with a significantly lower affinity (Mason et al., 1991). Ca²⁺ release, on the other hand, depends mainly on two types of Ca²⁺ release channels named inositol 1,4,5-trisphosphate (InsP₃) and ryanodine receptors (for reviews see Mikoshiba, 1993; Sorrentino and Volpe, 1993; Ehrlich, 1995). These channels are expressed in variable proportions in different cell types and couple extracellular stimuli to the release of Ca²⁺, with possible ensuing generation of Ca²⁺ waves and spikes (for reviews see Amundson and Clapham, 1993; Taylor, 1994; Bootman and Berridge, 1995). The relationship between these types of Ca²⁺-release channels is still largely debated. The ryanodine-sensitive channel is also activated by caffeine, and ryanodine- and caffeine-sensitive stores are generally regarded to comprise the same pool (Zacchetti et al., 1991; Barry and Cheek, 1994; but also see Giannini et al., 1992; McNulty and Taylor, 1993).In the vast majority of cell types so far investigated, the InsP₃- (and/or the ryanodine-) sensitive stores almost completely overlap with those sensitive to Tg (Zacchetti et al., 1991; Gamberucci et al., 1995) and are thus referred to also as Tg-sensitive Ca²⁺ pools. From the cytological point of view, the InsP₃-/Tg-sensitive Ca²⁺ pool is identified with the ER or with a subfraction of it (Hashimoto et al., 1988).The complexity of the relationships between the InsP₃- and ryanodine/caffeine-sensitive stores does not cover the entire issue of intracellular Ca²⁺ pool heterogeneity. Other types of Ca²⁺ pools are known to exist, the size of which varies considerably among different cell types. These latter Ca²⁺ stores account for roughly half of all sequestered Ca²⁺ (Chandra et al., 1991; Fasolato et al., 1991; Shorte et al., 1991; Bastianutto et al., 1995; Mery et al., 1996). They have been identified through the increase in [Ca²⁺]_i upon application of Ca²⁺ ionophores, after depletion of the Tgsensitive pool with a combination, or a sequence, of InsP₃generating agonists, Tg, and caffeine. These residual Tginsensitive pools appear rather heterogeneous in terms of cytological identity and pharmacological sensitivity. Part of these pools shows an acidic lumenal pH and is discharged only by a combination of a Ca²⁺ ionophore and of agents that collapse internal acidic pH gradients (such as monensin and NH₄Cl). ⁴⁵Ca²⁺ labeling of Tg-insensitive pools is slower than that of the Tg-sensitive store, and, for this reason, they have been generally indicated as slowly exchanging Ca²⁺ pools (Fasolato et al., 1991). As far as their identification is concerned, the acidic pool seems largely identifiable with secretory compartments and lysosomes, while very little is known yet about the rest of the Tg-insensitive store.Here we demonstrate that a nonacidic, InsP₃- and Tg- insensitive Ca²⁺ pool rapidly accumulates large amounts of Ca²⁺ when high and sustained increases of [Ca²⁺]_i are induced by opening of voltage- or store-operated Ca²⁺ channels. This Ca²⁺ storage compartment is insensitive to mitochondrial uncouplers and appears diffusely distributed in the cell cytosol. The possibility is discussed that this low-affinity, high-capacity Ca²⁺ pool represents a previously unidentified subcompartment of the ER expressing a Tg-insensitive Ca²⁺–ATPase.     

Evidence for a Slow-Turnover Form of the Ca2+-Independent Phosphoenolpyruvate Carboxylase Kinase in the Aleurone-Endosperm Tissue of Germinating Barley Seeds

Phosphoenolpyruvate carboxylase (PEPC) activity was detected in aleurone-endosperm extracts of barley (Hordeum vulgare) seeds during germination, and specific anti-sorghum (Sorghum bicolor) C₄ PEPC polyclonal antibodies immunodecorated constitutive 103-kD and inducible 108-kD PEPC polypeptides in western analysis. The 103- and 108-kD polypeptides were radiolabeled in situ after imbibition for up to 1.5 d in ³²P-labeled inorganic phosphate. In vitro phosphorylation by a Ca²⁺-independent PEPC protein kinase (PK) in crude extracts enhanced the enzyme''s velocity and decreased its sensitivity to l-malate at suboptimal pH and [PEP]. Isolated aleurone cell protoplasts contained both phosphorylated PEPC and a Ca²⁺-independent PEPC-PK that was partially purified by affinity chromatography on blue dextran-agarose. This PK activity was present in dry seeds, and PEPC phosphorylation in situ during imbibition was not affected by the cytosolic protein-synthesis inhibitor cycloheximide, by weak acids, or by various pharmacological reagents that had proven to be effective blockers of the light signal transduction chain and PEPC phosphorylation in C₄ mesophyll protoplasts. These collective data support the hypothesis that this Ca²⁺-independent PEPC-PK was formed during maturation of barley seeds and that its presumed underlying signaling elements were no longer operative during germination.Higher-plant PEPC (EC 4.1.1.31) is subject to in vivo phosphorylation of a regulatory Ser located in the N-terminal domain of the protein. In vitro phosphorylation by a Ca²⁺-independent, low-molecular-mass (30–39 kD) PEPC-PK modulates PEPC regulation interactively by opposing metabolite effectors (e.g. allosteric activation by Glc-6-P and feedback inhibition by l-malate; Andreo et al., 1987), decreasing significantly the extent of malate inhibition of the leaf enzyme (Carter et al., 1991; Chollet et al., 1996; Vidal et al., 1996; Vidal and Chollet, 1997). These metabolites control the rate of phosphorylation of PEPC via an indirect target-protein effect (Wang and Chollet, 1993; Echevarría et al., 1994; Vidal and Chollet, 1997).Several lines of evidence support the view that this protein-Ser/Thr kinase is the physiologically relevant PEPC-PK (Li and Chollet, 1993; Chollet et al., 1996; Vidal et al., 1996; Vidal and Chollet, 1997). The presence and inducible nature of leaf PEPC-PK have been established further in various C₃, C₄, and CAM plant species (Chollet et al., 1996). In all cases, CHX proved to be a potent inhibitor of this up-regulation process so that apparent changes in the turnover rate of PEPC-PK itself or another, as yet unknown, protein factor were invoked to account for this observation (Carter et al., 1991; Jiao et al., 1991; Chollet et al., 1996). Consistent with this proposal are recent findings about PEPC-PK from leaves of C₃, C₄, and CAM plants that determined activity levels of the enzyme to depend on changes in the level of the corresponding translatable mRNA (Hartwell et al., 1996).Using a cellular approach we previously showed in sorghum (Sorghum bicolor) and hairy crabgrass (Digitaria sanguinalis) that PEPC-PK is up-regulated in C₄ mesophyll cell protoplasts following illumination in the presence of a weak base (NH₄Cl or methylamine; Pierre et al., 1992; Giglioli-Guivarc''h et al., 1996), with a time course (1–2 h) similar to that of the intact, illuminated sorghum (Bakrim et al., 1992) or maize leaf (Echevarría et al., 1990). This light- and weak-base-dependent process via a complex transduction chain is likely to involve sequentially an increase in pHc, inositol trisphosphate-gated Ca²⁺ channels of the tonoplast, an increase in cytosolic Ca²⁺, a Ca²⁺-dependent PK, and PEPC-PK.Considerably less is known about the up-regulation of PEPC-PK and PEPC phosphorylation in nongreen tissues. A sorghum root PEPC-PK purified on BDA was shown to phosphorylate in vitro both recombinant C₄ PEPC and the root C₃-like isoform, thereby decreasing the enzyme''s malate sensitivity (Pacquit et al., 1993). PEPC from soybean root nodules was phosphorylated in vitro and in vivo by an endogenous PK (Schuller and Werner, 1993; Zhang et al., 1995; Zhang and Chollet, 1997). A Ca²⁺-independent nodule PEPC-PK containing two active polypeptides (32–37 kD) catalyzed the incorporation of phosphate on a Ser residue of the target enzyme and was modulated by photosynthate transported from the shoots (Zhang and Chollet, 1997). Regulatory seryl phosphorylation of a heterotetrameric (α₂β₂) banana fruit PEPC by a copurifying, Ca²⁺-independent PEPC-PK was shown to occur in vitro (Law and Plaxton, 1997). Although phosphorylation was also detected in vivo and found to concern primarily the α-subunit, PEPC exists mainly in the dephosphorylated form in preclimacteric, climacteric, and postclimacteric fruit.In a previous study we showed that PEPC undergoes regulatory phosphorylation in aleurone-endosperm tissue during germination of wheat seeds (Osuna et al., 1996). Here we report on PEPC and the requisite PEPC-PK in germinating barley (Hordeum vulgare) seeds. PEPC was highly phosphorylated by a Ca²⁺-independent Ser/Thr PEPC-PK similar to that found in other plant systems studied previously (Chollet et al., 1996); however, the PK was already present in the dry seed and its activity did not require protein synthesis during imbibition.     

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.