Plant Molecular Biology 38: 49–76, 1998. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

49

The molecular characterization of transport vesicles David G. Robinson∗, Giselbert Hinz and Susanne E.H. Holstein Abteilung Strukturelle Zellphysiologie, Albrecht-von-Haller Institut für Pflanzen-wissenschaften, Universität Göttingen, Untere Karspüle 2, 37073 Göttingen, Germany (∗ author for correspondence; e-mail [email protected])

Key words: clathrin-coated vesicles, COP-coated vesicles, dense vesicles, endocytosis, vacuolar protein targeting, v/t-SNAREs Abstract Secretion, endocytosis and transport to the lytic compartment are fundamental, highly coordinated features of the eukaryotic cell. These intracellular transport processes are facilitated by vesicles, many of which are small (100 nm or less in diameter) and ‘coated’ on their cytoplasmic surface. Research into the structure of the coat proteins and how they interact with the components of the vesicle membrane to ensure the selective packaging of the cargo molecules and their correct targeting, has been quite extensive in mammalian and yeast cell biology. By contrast, our knowledge of the corresponding types of transport vesicles in plant cells is limited. Nevertheless, the available data indicate that a considerable homology between plant and non-plant coat polypeptides exists, and it is also suggestive of a certain similarity in the mechanisms underlying targeting in all eukaryotes. In this article we shall concentrate on three major types of transport vesicles: clathrin-coated vesicles, COP-coated vesicles, and ‘dense’ vesicles, the latter of which are responsible for the transport of vacuolar storage proteins in maturing legume cotyledons. For each we will summarize the current literature on animal and yeast cells, and then present the relevant data derived from work on plant cells. In addition, we briefly review the evidence in support of the ‘SNARE’ hypothesis, which explains how vesicles find and fuse with their target membrane.

Introduction The transfer of transmembrane proteins and lumenal macromolecules (proteins, glycoproteins, and polysaccharides) between organelles of the endomembrane system (endoplasmic reticulum (ER), Golgi apparatus (Gapp), endosomes, lysosomes and vacuoles), and between the plasma membrane (PM) and these compartments, takes place by vesicle transport. Many, if not all, of the vesicles involved are formed with a protein covering ‘coat’ on their cytoplasmic surface, which has later to be removed to allow for vesicle fusion. Vesicle transport is therefore a multistep process: it starts by the sequestration of cargo molecules at a particular locus in the membrane of the donor organelle. This site is usually predetermined by the attachment of coat proteins, but their recruitment may also occur simultaneously or subsequent to

the collection of the cargo molecules. Vesicle formation and budding are a consequence of the continued assembly of the coat proteins, for which both ATP and a small GTPase are normally required. Vesicle targeting, on the other hand, appears to involve the recognition of proteins specific for the vesicle membrane (so-called v-SNAREs) by proteins specific for the target organelle (t-SNAREs). Correct docking may or may not be preceded by the dissociation of the coat proteins, an event also needing ATP and/or GTP. The ultimate act of fusion may be less understood, although it is clear that, at least for exocytosis, Ca2+ ions play a major role in this process (see Thiel and Battey, this volume). Work on non-plant organisms has established that there are a minimum of five coated vesicle types: two, and possibly a third, contain clathrin together with adaptor complexes (AP). The others, called COP vesi-

50 cles, have a coat comprising a multimeric ‘coatomer’ and a GTP-binding protein. These have been extensively investigated and a number of excellent reviews are available, most recently by Schmid [214] for clathrin-coated-vesicles (CCV), and Rothman and Wieland [202], Schekman and Orci [211] for COP vesicles. Plants also have CCV, which were first isolated in 1982 [139], but progress on their biochemical characterization has been slow, in the main due to poor recoveries and inadequate protection against proteolytic degradation during their isolation. Two major reviews on plant CCV appeared in 1996 [16, 190]. In the meantime, several groups are now investing in a molecular biological approach which is already beginning to pay dividends. Although the object of much speculation (see, for example, the latest major review on the plant Gapp [235]), COP vesicles remain to be purified and characterized from plant extracts. However, as we shall describe later, the tools are now becoming available with which it should be possible to alleviate this situation in the very near future. In addition to the above types of transport vesicles, there are several different types of Golgi-derived vesicles which, in thin section, appear to be smoothsurfaced. Some of them, including the immature secretory granules of exocrine glands and the ‘dense vesicles’ (DV) in developing seeds, contain condensed protein aggregates. Others, such as the slimecontaining vesicles of root cap cells, have less structured contents. Interestingly, CCV are often to be seen budding from the surface of both types of vesicles, an observation frequently interpreted as representing a retrieval mechanism for missorted proteins (see below). Although secretory vesicle-containing fractions from plants have been separated from plant homogenates [187, 240], their purity is questionable and no attempts at analysing their membranes have been undertaken. By contrast, DV, which transport proteins to the storage vacuole, have recently been successfully isolated and subjected to a preliminary characterization [87].

late endosomes, where they recycle receptors back to the cell surface [239]. Although CCV have not been visualized in yeast cells in situ, disruption of the clathrin heavy- and light-chain genes (CHC1, CLC1) as well as the production of temperature-sensitive CHC1 mutants have resulted in cells with an impaired ability to perform receptor-mediated endocytosis of the pheromone α-factor and which are also defective in the sorting of soluble vacuolar proteins [31, 93, 165, 222]. Moreover, in clathrin-disrupted cells the late Golgi-localized endoprotease Kex2p is no longer retained and is found instead at the cell surface [188]. Clathrin-deficient mutants in Dictyostelium discoideum also have severely reduced endocytotic activity, and are unable to form contractile vacuoles. In addition, they show aberrant cytokinesis [152, 155]. In plants CCV are also seen to be formed at the PM and the TGN (Figure 1), but unlike mammalian and yeast cells there is no unequivocal proof as yet for receptor-mediated endocytosis, although the uptake of unspecific electron opaque tracers via CCV has been demonstrated on numerous occasions (see Griffing, this volume). Thus, the possibility that PM-derived CCV merely serve to retrieve membrane from the cell surface cannot be ruled out [22, 206]. Multivesicular bodies, which are endosomal in character [193, see also Griffing, this volume), are quite often seen with a partial coat (see, for example, Figure 8a in [191]), but it is unclear whether this represents the start of vesiculation, nor has it been shown unequivocally that clathrin is present in the coat. For a number of years TGN-derived CCV in plants were considered to contain vacuolar storage proteins (reviewed in [16]). A meticulous analysis of subcellular fractions, together with in situ immunogold labelling [88], however, has refuted this notion. On the other hand, the currently favoured hypothesis that TGN-derived CCV are involved in the transport of acid hydrolases still requires final proof (see below). Clathrin

Clathrin-coated vesicles Sites of formation and general functions In mammalian cells CCV have been shown to bud from three different membranes: the PM, where they function to internalize cell surface receptor-bound ligands [198]; the trans-Golgi network (TGN), where they are responsible for the selective transport of lysosomal acid hydrolases out of the Gapp [25]; and

Clathrin triskelions constitute the outermost, cytoplasmic-facing layer of a CCV (Figure 2A), and all CCV derive their triskelions from the same cytoplasmic pool [65]. Triskelions are hexameric, with three heavychain (CHC) and three light-chain (CLC) polypeptides. CHC have the intrinsic property to spontaneously assemble under defined conditions (low-ionicstrength buffer, 2 mM CaCl2 and low pH) into a heterogeneous population of closed polyhedral struc-

51

Figure 1. Clathrin-coated vesicles. a. Formation at the plasma membrane of the growing cell plate in suspension-cultured tobacco BY-2 cells. Cross-sectioned pits (arrows), as well as planar clathrin lattices (double-headed arrows) are visible in this tangential section. b, c. Formation at the plasma membrane of an expanding cell in cauliflower inflorescence. Invagination (coated pit, b) and constriction (release of coated vesicle, c) are readily recognized. d. Formation at the trans-Golgi in suspension-cultured carrot cells. Arrows point to budding profiles of coated vesicles, some distance away from the stack. e. Negatively stained clathrin-coated vesicles isolated from developing pea cotyledons. Empty ‘cages’ are also visible (large arrow) which show clearly the typical polygonal arrangement of the clathrin triskelions. Bars = 100 nm.

52 tures called ‘cages’. Under physiological conditions, however, clathrin assembly is dependent upon the presence of AP (see below). In their completed form cages have the structure of a fullerene, i.e. the vertices have three ordinates and all of the faces are either pentagons or hexagons. To form a closed shell 12 pentagons must always be present, but according to the size of the cage the number of hexagons may vary (for pertinent literature, see [180]). Rotary shadowed preparations of triskelions in the electron microscope reveal their bent, three-legged structure. These structures are, however, non-planar and puckered at their vertices [107]. Each leg is 45 nm in length in mammalian triskelions, and three morphologically distinct regions can be recognized: a protease-sensitive knee, separating 17 nm proximal (adjacent to the vertex) and 25 nm distal domains (Figure 2D). The latter terminates in a 52–59 kDa globular N-terminal domain, which appears to be connected to the distal portion of the leg via a flexible 6.5– 10 nm linker region [105, 252]. Triskelions from yeast and plants are morphologically identical to those from mammalian CCV, but, consistent with their higher molecular masses, have total leg lengths of 61 nm [34]. CHC polypeptides are highly conserved and are encoded by a single gene in all eukaryotes. The 1675 amino acid (aa) CHC sequence from rat codes for a 191 kDa polypeptide, which runs in SDS-PAGE at 180 kDa. Mammalian CHC share 99% identity with each other and 50% and 57% similarity to yeast and Dictyostelium respectively (see [22] for literature). Sequence comparison between the 1700 aa sequence of soybean (Glycine max) with rat and bovine CHC sequences reveals a remarkably high degree of conservation: 56% identity to rat, 54% to Dictyostelium, and 44% to yeast, with the greatest degree of divergence confined to the N- and C-termini. Nevertheless, with the exception of one report [33], there seems to be little or no recognition of clathrin by heterologous CHC antisera [50]. Controlled proteolysis of mammalian CHC as monitored by high-resolution electron microscopy has shown that the distal segment contributes to the stability of the clathrin lattice, while the hub region (proximal leg domain and vertex, residues 1074–1675) is responsible for CHC trimerization, CLC binding, and cage assembly. The trimerization domain has been narrowed down to residues 1550–1615, which includes a region of α-helical conformation. The remaining 70 C-terminal residues, rich in proline and glycine and absent in yeast CHC, which protrude from

the top of the vertex, are not required for trimerization (see [124] for literature). Residues 1438–1481 comprise the CLC-binding region, with an overlapping stretch of amino acids (residues 1460–1489) which is predicted to form a coiled-coil helix [144]. Current models for CHC/CLC interaction envisage a four helix bundle with both the CLC and CHC folding back on themselves and the C-terminus of the CLC pointing towards the vertex [106, 124]. In contrast to these data, yeast CLC are described as binding directly to the trimerization domain, acting as a hinge between this and the proximal domain of the CHC [178]. Adaptorbinding sites on the CHC have also been identified: one at the N-terminus, projecting inwards towards the vesicle membrane surface, and the other in the hub region [143, 100]. In mammalian cells two distinct, single-copy genes for the light chains have been identified: one encoding for CLCa , with a predicted molecular mass of 27.5 kDa (33 kDa in SDS-PAGE), and the other for CLCb (27 and 36 kDa, respectively). Yeast has only one CLC with a molecular mass of 26.5 kDa (predicted) and 38 kDa (in SDS-PAGE). While the amino acid sequences of the two mammalian CLCs are 60% identical to one another [96], the single yeast CLC shares only 18% sequence similarity with its mammalian counterparts [226]. The diversity of mammalian CLC is also a consequence of tissue-specific mRNA splicing in neurons [236]. Candidates for CLC from plants have been described: two polypeptides with molecular masses in SDS-PAGE of 45 and 52 kDa from zucchini hypocotyl CCV [44], and several polypeptides from pea cotyledon CCV with molecular masses ranging from 40 to 50 kDa [121]. The situation will remain unclear until sequence data become available. Adaptor complexes AP were originally identified as coat protein fractions from bovine brain CCV which could be separated from clathrin by gelfiltration and which promoted the assembly of clathrin into cages under physiological conditions in vitro. Further purification by hydroxyapatite chromatography (HA) led to the identification of the two complexes HA-1 and HA-2, which were later renamed AP-1 and AP-2 [99]. The more recent term, adaptor complex, reflects more suitably their function in interacting with sorting signals in the cytoplasmic domains of transmembrane receptors on the one hand

53

Figure 2. Diagrammatic representation of the various coat polypeptides and vesicle targeting. A. Clathrin-coated vesicle. B. COP-I vesicle. C. COP-II vesicle. D. Clathrin triskelion. E. Adaptor complexes: AP-1 (β1, γ , µ1, σ 1); AP-2 (α, β2, µ2, σ 2). F. Components of the targeting machinery, and steps in vesicle docking and fusion.

54 [156], and triskelion binding to membranes [244] and membrane recognition on the other [131, 169]. Mammalian adaptors Three different but homologous AP have been described for mammalian cells: AP-1, which is localized to the TGN, early endosomes and immature secretory granules [6, 47, 114], AP-2, mainly present at the PM [168] but also detected on endosomes and lysosomes [221, 245], and AP-3 found on the TGN/and or endosomes, which, in contrast to AP-1 and AP-2, is not associated with clathrin [43]. All of these AP are heterotetrameric. The constituent polypeptides, named adaptins, fall into three categories: large adaptins: α, β1, β2, β3, γ at around 100 kDa (δ at 130 kDa); medium adaptins: µ1, µ2, µ3 at about 50 kDa; small adaptins: σ 1, σ 2, σ 3 at about 20 kDa [6, 227]. A certain degree of homology exists between the adaptins of each group (see Table 1). Each complex contains two large subunits, and one each of the medium and small subunits (see Figure 2E). Electron microscopy has shown AP-2 to have a three-domain structure: a brick-shaped core, containing the N-terminal bulk of the adaptins to which the medium and small chains are attached, connected via a flexible hinge to an ear-like appendage [79]. Trypsin primarily attacks the hinge leaving a proteolysis-resistant 60–70 kDa truncated adaptor; the AP-1 adaptor behaves similarly [217]. With the exception of the σ -adaptins, the function(s) of the adaptins have now been established and the responsible structural domains localized (summarized in Table 2). Clathrin binding is a property of the β-adaptins, which need to be intact for highaffinity interaction [217]. They alone can drive the in vitro assembly of clathrin into cages [59]. Although there is evidence that the core region of the αA -adaptin can bind to clathrin it fails to stimulate clathrin assembly [64]. The α- and γ -adaptins, rather than the β-subunits, are responsible for membrane attachment with the binding domain residing in the Nterminal core [29, 162, 197]. Based on experiments in which the recruitment of chimeric adaptors (in which the trunk portions of the α- and γ -adaptins were exchanged) was investigated, it would appear that membrane selection is also a property of the µ- and σ adaptins [162]. Thus, µ1/σ 1-adaptins direct adaptors to the TGN, and adaptors with µ2/σ 2-adaptins bind to the PM. Furthermore, the µ-adaptins show a strong affinity for both β-adaptins, while the σ 1- and the σ 2-adaptins bind to the N-terminus of the γ - and α-

subunits, respectively, suggesting that they may play a regulatory role in AP assembly and membrane recruitment. In terms of interactions with other proteins α-adaptin is by far the most versatile, with binding sites for clathrin, dynamin, eps15, inositol phosphates and synaptotagmin (see Table 2). All adaptins, except for the σ -subunits, can be phosphorylated (in vitro: α, β1, β2, γ , µ2 [140]; in vivo: α, β1, β2, µ1, µ2 [260]). Phosphorylation of the large adaptins in vivo is restricted to the hinge region, and the phosphorylated forms are found predominantly in the cytosol, implicating a regulatory role for dephosphorylation in membrane recruitment. Adaptins in non-mammalian systems Adaptin homologues have been identified in a number of other eukaryotes, including plants. α-adaptins from the nematode Caenorrabditis elegans and Drosophila possess 64% and 60% respectively, overall identities to mammalian αA -subunits [49, 261]. A β-homologue has been described from Drosophila, which in its N-terminal domain (residues 1-575) is a functional hybrid of the mammalian β1- and β2-adaptins, and therefore colocalizing with both the α-adaptin at the PM and with the γ -adaptin at the TGN [27]. In yeast two sequences homologous to β-adaptins, one with a 35% identity to β2, have been reported [103, 185], but the proteins remain to be isolated. A β-homologue in plants has been identified using monoclonal antibodies directed against mammalian β-adaptins [89]. This adaptin responded in the same way to tryptic digestion as its mammalian counterparts, and behaved biochemically like a β1-adaptin in HA chromatography although it was localized to the PM [50]. γ -adaptin homologues from the basidiomycete Ustilago maydis and from Arabidopsis, each with 48% identity to mouse γ -adaptin have been reported [101, 213]. Two µ-subunit homologues have been described from C. elegans, displaying 47% and 42% overall identities to the µ1- and µ2-adaptins of mouse, respectively [115]. µ-type adaptins have also been identified in the slime mould Dictyostelium (52% identity to µ2 of rat [242]), in yeast (APM1 with 56% identity to mouse µ1 [146]; APM2 with 39% identity to mouse µ2 [238]), and in plants (with 48% a slightly higher identity to the µ2 of C. elegans than to the µ1 of C. elegans or mouse with 41–44% [68]). The µ1adaptin of bovine brain has a serine kinase exhibiting casein kinase II-like properties [138]. A similar kinase

55 Table 1. Adaptor complexes and their homologues.

Large subunits (ca. 100 kDa)

AP-1 CCV

AP-2 CCV

AP-3

γ

αA , αC

δ

COP-I homologues

Plant homologues γ

/ β1

β2

β3A /β3B

β-COP

β2

Medium subunit (ca. 50 kDa)

µ1

µ2

µ3A , µ3B

δ-COP

µ

Small subunit (ca. 20 kDa)

σ1

σ2

σ 3A , σ 3B

ζ -COP

σ 1, σ 2

Homology [reference] αA , αC : 84% [195] γ /α: 25% [196] α, γ /δ: 25%1 [158] β1/β2: 84% [181] β1; β2 / β-COP: 17%1 [52] β3A /β3B : 61% [42] µ1/µ2: 40% [146] µ1; µ2/µ3A /µ3B : 27–30% [174] µ3A /µ3B : 80% [174] µ1; µ2/δ-COP: 22–24%(1 [55] σ 1/σ 2: 45% [108] σ 1; σ 2/σ 3A ;σ 3B : 29–31% [43] σ 3A / σ 3B : 84% [43] σ 1; σ 2/ζ -COP: 19–29% [37]

1 N-terminal homology. 2 Immunological evidence.

Table 2. Functions of adaptor subunits. AP-subunit

Functional regions

Function [reference]

αA (108 kDa) / αC (104 kDa)

n.d. N-terminal: 5–80 132–331 core 29 kDa αA core fragment hinge C-terminal 40 kDa fragment ear region C-terminal 701–938

Binding of synaptotagmin [265] Inositol polyphosphate binding [58] PM-targeting/σ 2 subunit interaction [162] self association [13] Clathrin binding [64] Phosphorylation site [260] eps 15 interaction [18] PM-targeting [162] Dynamin-binding [253]

β1 (115 kDa) / β2 (106 kDa)

N-terminus

ASG-R interaction [17]

core

Armadillo-repeats: prot./prot.-interaction [111] µ chain binding site [162] Phosphorylation site [262] Clathrin-binding site [224]

hinge hinge: 616–663 γ

N-terminal 132–331 res. 566–594

µ1 (47 kDa) µ2 (50 kDa)

N-terminal 1–145 C-terminal 147–423 N-terminal 1–145 C-terminal 164–435

β-adaptin interaction [3] Binding to YXXØ motifs [3] β-adaptin interaction [3] Binding to YXXØ motifs [3] Membrane selection? [162]

n.d. n.d.

Membrane selection? [162] Membrane selection? [162]

µ1/µ2 σ 1 (19 kDa) σ 2 (17 kDa) n.d.: not determined.

TGN-targeting [162] σ 1/µ1-chain interactions [162] β1-chain binding

56 activity has been described for an adaptor-containing fraction from zucchini hypocotyl CCV [51]. Homologues to mammalian σ -adaptins have been identified in yeast (APS1 with 53% identity to mouse σ 1 [108]; APS2 with 50% identity to mouse σ 2 [145]), and plants (a σ 1 homologue in chinese medical tree with 70–80% similarity to mouse σ 1 and a 65–73% similarity to σ 2 of rat and yeast [129]; a σ 2 homologue from maize with 65% similarity to σ adaptins of rat and man [199]). Thus, while intact AP have not yet been isolated from non-mammalian systems, the identification of homologues for nearly all of the well-characterized adaptins from mammals points heavily towards their existence in the same tetrameric form in these other organisms. Other clathrin-binding proteins Neuronal tissue contains two other proteins which are minor components of CCV, auxilin and AP180 [4, 5]. Both are monomeric proteins of about 90 kDa and both act as cofactors for clathrin assembly in vitro under physiological conditions, with AP180 being about four times as effective as the adaptors or auxilin in this regard [122]. The sequence and proteolytic cleavage pattern of AP180 suggests a three domain structure, with the N-terminal 300 residues (33 kDa) comprising a clathrin-binding region. The acidic middle portion appears to be responsible for the anomalous physical properties of the protein (molecular mass in SDS-PAGE 155–180 kDa; predicted size 91.4 kDa [141]). The 33 kDa AP180 fragment binds inositol polyphosphates with high affinity thus preventing cage assembly in vitro [262]. Auxilin shows significant homology in its N-terminal half (residues 47–350) to the actin-binding protein tensin [218] and an analysis of its complete structure reveals it to be a DnaJ-like protein, and thus the partner for the uncoating ATPase Hsc70 (see below). Homologues for these proteins in other organisms have not yet been discovered. CCV formation Receptor-coat protein interactions Mammalian cells. The PM, TGN, and endosomes are the three main sites for receptor-mediated clathrincoupled sorting events in animal cells [132, 214, 246]. Some of the receptors at the PM of mammalian cells are always found concentrated in coated pits, and are constitutively internalized via CCV, irrespective of

whether cargo molecules (ligands) are bound or not. Examples of this type of receptor are the low-density lipoprotein receptor (LDL-R), the transferrin receptor (Tf-R), and the cationic-independent mannose-6phosphate receptor (ci-MP-R). Another class of receptors, including the epidermal growth factor receptor (EGF-R), is first concentrated in coated pits after successful ligand binding, a process which also triggers a tyrosine kinase activity contained in the cytosolic domain of the receptor. For both types of receptor an interaction with AP-2 has been established [63, 167, 231]. The first destination of PM-derived CCV is the early endosome where receptor-ligand uncoupling occurs. Recycling of many of these receptors to the PM then follows by a separate set of CCV, which are smaller than those derived from the PM. The nature of the endosomal AP remains to be determined, but they contain neither α- nor γ -adaptins [239]. At the TGN newly synthesized acid hydrolases are specifically diverted from the secretory pathway through the participation of mannose 6-phosphate receptors: the 300 kDa ci-MP-R and the 45 kDa cationdependent MP-R (cd-MP-R) (reviewed by [25]). An interaction between AP-1 and the cytoplasmic tails of these receptors was demonstrated some years ago (e.g. Mauxion et al. [133]). MP-R-ligand complexes are then transported via CCV to a prelysosomal, endosomal compartment from which the receptors are recycled, again via CCV to the TGN. Both types of MP-R are found in addition at the PM, but only the ciMP-R is able to bind ligands (e.g. missorted, secreted lysosomal enzymes). Essentially two types of sorting signals exist in the cytosolic tails of the transmembrane receptors just mentioned: a tyrosine or phenylalanine-containing motif (YXRF in the TF-R; NPXY in LDL-R; YSKV and YKYSKV in ci-MP-R; YRGV and FPHLAF in cd-MP-R) and a dileucine motif (LLHV in the ci-MPR; HLLPM in cd-MP-R). Recognition of both types of signal by AP-1 and AP-2 has been unequivocally demonstrated [75, 132, 157]. While the binding of the tyrosine-based motif occurs via the µ1- and µ2adaptins, the binding partner for the dileucine motif within the AP complexes remains to be elucidated [156]. In general the tyrosine-containing motifs mediate internalization at the PM, but only a subset is involved in lysosomal targeting. Thus, AP-2 are considered to have either a broader specificity or a higher overall affinity for sorting signals relative to AP-1. In analogy to the µ-adaptins of AP-1 and AP-2, the µ3-adaptin of AP-3 has recently been shown to recog-

57 nize the sequence YQRL in the cytoplasmic tail of the protein TGN38 [43]. As with many transmembrane receptors, both MPR contain multiple sorting signals in their cytoplasmic tails, which are responsible for their correct targeting to different compartments. Thus, for endocytosis at the PM the ci-MP-R requires only a single motif (YSKV), whereas for the cd-MP-R two motifs are necessary (YRGV and FPHLAF, the latter dominating). However, for lysosomal enzyme sorting the reverse is true: the ci-MP-R needs two signal motifs (LLHV and YKYSKV), whereas the dileucine motif HLLPM is sufficient for the cd-MP-R. In both MPRs the dileucine motifs are flanked by casein-kinase phosphorylation sites (ESEER sequence in the cd-MPR), which are phosphorylated in vivo, and which are necessary for the high-affinity binding of the AP-1 complexes [133, 138]. Clathrin or specific phosphoinositides in the PM may also increase the affinity of the µ2-adaptin for tyrosine-based endocytic motifs [186]. Yeast and plants. Binding of the α-factor by the yeast PM receptor Ste2p resembles that of the Gprotein-coupled receptors in inducing signal transduction. However, in contrast to the animal receptors this requires prior ubiquitinylation of the receptor [81]. In the internalization sequence SINNDAKSS mutation of the lysine as well as of the three nearest serine residues abolishes both ubiquitinylation and internalization. Phosphorylation of Ste2p is also required for its uptake. Kex2p, an integral membrane protein of the late Golgi compartment in yeast, is responsible for the proteolytic processing of α-factor; it possesses a tyrosine motif in its cytoplasmic tail [259]. Loss of clathrin function results in the delivery of Kex2p to the cell surface [166]. Similarly, the cytoplasmic tails of Vps10p, the receptor for vacuole-destined carboxypeptidase Y, and the membrane-bound proform of alkaline phosphatase, also a vacuolar enzyme, contain tyrosine motifs [38, 234]. The former is considered to interact with AP-1, the latter with AP-3-like adaptors, whereby AP-3-mediated vacuolar traffic appears to bypass the endosomal/prevacuolar compartment in yeast [39]. In plants the vacuolar protein sorting receptor BP80 (AtELP) is enriched in CCV [7, 87, 110, 193, see also Müntz; Neuhaus and Rogers, this volume]. Its structure resembles that of EGF in the lumenal, extracytoplasmic domain with several cysteine-rich domains, while its cytoplasmic domain contains two

tyrosine-based sorting signals (residues 589–594 and 606–609), with only the latter one fitting the consensus motif Yxxφ [7, 163]. Experiments have recently been performed by Beevers and colleagues [26] which suggest that BP-80 from pea cotyledons can bind adaptors from bovine brain, wheatgerm and pea cotyledons. It has been claimed that this binding is localized to a tyrosine-containing motif in the cytoplasmic domain of BP-80. Heterologous binding experiments have also been carried out in our own laboratory [194]. These have involved the cytoplasmic domain of the ci-MPR and CCV coat proteins from zucchini hypocotyls. When zucchini cages (containing adaptors) are incubated together with the MP-R tail, the latter is quantitatively bound; other non-receptor proteins did not associate with the cages. These experiments point to a similarity in adaptor-receptor binding mechanisms between plants and animals. Recruitment of adaptors and triskelions Although, as just described, AP do interact with sorting signals in the cytoplasmic tails of transmembrane receptors and are essential components of the adaptor docking site, these interactions alone are not sufficient to explain the observed tight membrane binding of adaptors. Our knowledge of the mechanisms underlying the assembly of CCV coat proteins onto membranes, especially the PM is mainly due to the use of cell-free and semi-permeabilized mammalian cell systems which have yet to be established in the field of vesicle-mediated protein transport in plants. This work has been excellently reviewed elsewhere [214, 215] so only a brief summary is necessary here. PM-derived CCV. Six stages are envisaged in the formation of CCV at the PM: AP-2 recruitment, assembly of clathrin into a planar lattice, dynamin recruitment, invagination to form a coated pit, constriction of the neck of the coated pit through dynamin rearrangement, and CCV release. There is an ongoing search for AP-2 binding proteins in the PM: a good candidate in neuronal tissue is synaptotagmin, whose cytoplasmic tail is known to bind to AP-2 in vitro [265]. GTP, ATP and other cytosolic factors are required for AP-2 recruitment from cytosolic fractions in vitro. GTPγ S, but not brefeldin A (BFA), inhibits AP-2 binding to the PM [28] and leads instead to their attachment on late endosomes [220]. A possible candidate for the required GTPase is ARF6 (ADP ribosylation factor 6), whose action is BFA-insensitive and which is present at the PM, although not ap-

58 parently specifically localized to coated pits [173]. Most recently, AP-2 recruitment has been shown to be neomycin sensitive, demonstrating that phospholipase D (which is activated by ARF) is also involved in this process [256]. Bound AP-2 are a prerequisite for clathrin recruitment. Only cytosolic, and not extracted and purified clathrin, can serve this purpose in vitro. Depending on the assay used, a controversy exists about the requirement for nucleotide and/or cytosolic cofactors. The gradual inwards curvature leading to invagination is believed to be a consequence of the insertion of pentagonal rather than hexagonal triskelion units [104]. A characteristic of coated pits in mammalian cells is their relatively long neck. Concentrated in this constriction is a special 100 kDa GTPase known as dynamin. Not only does dynamin interact with AP-2 [253] but it has the property of self-assembling into helical stacks [85]. It is thought that GTP binding causes dynamin to be redistributed from the lattice to the neck region, and that GTP hydrolysis leads to a tightening of the dynamin helix. For the final act of CCV detachment ATP is also required but the ATPase responsible has not yet been identified.

TGN-derived CCV. For recruitment of AP-1 onto Golgi membranes only the core domain is sufficient for binding [244]. Putative AP-1 docking proteins in the TGN have been demonstrated using coimmunoprecipitation: three novel proteins, p75, p80 and p60, were found specifically cross-linked to γ -, β1- and µ1-adaptins, respectively [221]. By passing detergent extracts of Golgi fractions over immobilized AP-1 an 83/52 kDa dimeric protein has also been identified [131]. Binding was restricted to the 83 kDa partner. AP-1 has recently also been shown to co-localize with syntaxin 6 (see SNARE hypothesis, below) on TGN membranes [23]. AP-1 recruitment differs from that of AP-2 at the PM and endosomes in being not phospholipase D-dependent, but resembles AP-2 recruitment onto endosomes in that it is inhibited by BFA [256]. AP-3 recruitment at the TGN is also prevented by BFA [227]. AP-1 and AP-3 recruitment therefore resembles that of COP-I coatomers (see below) in that it is enhanced by GTPγ S, indicating the participation of ARF1. Until very recently dynamin was not considered to play a role in CCV formation at the TGN, but evidence for a dynamin-like candidate has now been published [77].

CCV assembly in non-mammalian cells. Currently there is no information available on the mechanisms underlying the recruitment of AP and triskelions in fungi or plants, but both ARF [137, 189] and dynamin [60, 164, 251] homologues have been described in these organisms. In contrast to mammalian cells the yeast dynamin homologue Dnm1p is involved in endosomal vesicle trafficking rather than CCV budding at the PM since its deletion does not affect internalization of the pheromone α-factor. Similarly, the other yeast dynamin homologue, Vps1p, plays an important role in vacuolar protein transport. The Arabidopsis dynamin homologue ADL1 is similar to mammalian dynamin 1, but its intracellular location and mode of action remain to be determined. Uncoating CCV must shed their coats in order to fuse with a target membrane. Early studies demonstrated that a 70 kDa protein from brain cytosol was capable of dissociating triskelions from CCV in vitro in an ATPdependent manner. This protein was subsequently identified as a heat shock protein and designated as the Hsc70 uncoating ATPase [248]. Deep-etch rotary shadowed preparations have revealed that three Hsc70 molecules can bind to the surface of a triskelion at its vertex [80]. Originally it was considered that the presence of both CLC was necessary for the initial interaction between the uncoating ATPase and the triskelion, and that the N-terminal globular domain of the CHC functioned as a second binding site for Hsc70 [216]. However, it is now known that neither of these are required for uncoating [249]. A 100 kDa protein cofactor has been shown to be required for the uncoating reaction when highly purified clathrin cages, which are stabilized by AP-2, are used as a substrate [182]. This is the DnaJ-like protein auxilin [249]. DnaJ-like proteins are known to cooperate with various members of the Hsc70 family in diverse functions such as protein folding, transport of proteins across membranes, and dissociation of protein complexes [40]. In a manner analogous to the DnaJ/DnaK reaction, auxilin first attaches to the clathrin lattice, priming the basket for subsequent recruitment of Hsc70, which is in its ATP status. Hydrolysis of ATP finally leads to the release of the triskelions. When expressed within a 38 kDa C-terminal fragment, the clathrin-binding domain (residues 574– 814) and the J-domain (residues 813–910) are alone sufficient for the uncoating reaction [90]. Equally,

59 a 60 kDa fragment of Hsc70, which contains the ATPase- and substrate-binding domains is sufficient to dissociate triskelions from CCV in vitro [250]. So far, auxilin has only been found in neuronal tissue [218]. Stable cytosolic complexes of clathrin/Hsc70, together with a third component (valosin containing protein [179] or p532 [200]) have been reported suggesting that these proteins might participate in the regulation of the status of assembled to disassembled clathrin. After clathrin dissociation, the AP are retained by the uncoated vesicles [80]. Recent studies speculate on a second ATPase for adaptor release [232] as well as a protein, p90, which enhances AP release [84]. The first evidence for a CCV uncoating ATPase from plants has been presented in a study by Kirsch and Beevers [109]. A 70 kDa uncoating ATPase was purified from pea cotyledon extracts. This ATPase was capable of uncoating both pea cotyledon and bovine brain CCV. COP-coated vesicles Discovery and location In the early 1980s, Rothman and colleagues introduced the use of a cell-free system to follow the process of intra-Golgi transport. The transfer of the vesicular stomatitis virus (VSV) coat G protein between Golgi stacks isolated from infected wildtype and glycosylation-mutant Chinese hamster ovary (CHO) cells served as their assay. It was shown that successful transport-coupled glycosylation required both the presence of cytosol and ATP (reviewed in [201]). When, however, the Golgi fractions were examined in the electron microscope it became immediately clear that the VSV-G protein was not present in CCV, but rather in another sort of coated vesicle [159]. It was then found that GTPγ -S blocked the in vitro transport of VSV-G protein, resulting in a severalfold accumulation of non-clathrin-coated vesicles at the surface of the Golgi cisternae [135]. This observation was put to use by Malhotra et al. [130] who, by high-salt treatment followed by density gradient centrifugation, were able to dissociate and separate the vesicles from the Golgi membranes. These steps are graphically depicted in Figure 3. Subsequent isolation of the protein complex from the cytosol [255] and the introduction of the terms ‘coatomer’ for the complex, and ‘COPs’ for the constituent polypeptides, led to the term COP-coated vesicle (Figure 2B, C).

Table 3. Coat components of COP vesicles. Protein in mammals

Protein in yeast

Molecular mass (kDa)

Ret1p Sec26p Sec27p Sec21p Ret2p

Small GTPase

α-COP β-COP β 0 -COP γ -COP δ-COP ε-COP ζ -COP ARF1

Ret3p ARF1p

160 110 102 98 61 31 20 20

COP-II Sec23 complex

hSec23A

Sec13 complex

Sec13Rp

Small GTPase

Sar1a/b

Sec23p Sec24p Sec13p Sec31p Sar1p

85 105 33 105 21

COP-I Coatomer

Roughly at the same time, work on yeast had identified genes whose products were found to be absolutely necessary for the successful transport of the pro-form of α-factor from the ER to the Golgi in vitro; amongst these were Sec12p, Sec13p, Sec16p, Sec 23 and Sar1p [98, 183]. This enabled Schekman’s group to isolate a second type of COP-coated vesicle, termed COP-II vesicles, by incubating nuclear envelopes (equivalent to ER) in the presence of ATP, GTP and three soluble proteins: Sar1p, Sec13p complex and the Sec23p complex [10]. In the meantime, ER-derived COP-II vesicles from mammalian cells have been identified in situ [160] and isolated [203]. COP-I coat components ARF1 ARF1 is a 20 kDa GTP-binding protein (see Table 3), which is found mainly in the cytosol in monomeric GDP form [223]. The GTP form is Nterminally myristoylated, allowing for membrane anchorage, which can occur in the absence of coatomers [48]. In comparison to the other ARFs [119] binding of ARF1 requires a GTP/GDP exchange factor (GEF), which itself is membrane-bound [76]. GEF is the target for BFA [48], which is known to block protein transport through the Gapp by causing the release of ARF1-attached coatomers [210].

60 Coatomers The coatomer is a 700 kDa heterooligomeric complex comprised of seven stoichiometric proteins (see Table 3). These proteins conveniently fall into three groups, analogous to the coat proteins of CCV: a large subunit (α-COP), medium subunits (β-, β 0 -, γ , δ-COPs), and small subunits (ε-, ζ -COPs). In fact, homologies (19–29% identities) between the medium and small coatomer subunits and the CCV adaptins do exist (see Table 1), although the two groups of coat proteins are immunologically distinct. Interestingly, so-called WD-40 motifs, which are often found in heterooligomeric protein complexes [147], are typical of α- and β 0 -COPs. Coatomer assembly in vivo has recently been studied [125], and shown to be a very coordinated process taking 1–2 h to complete. Direct interactions occur between α-, β 0 - and δ-COPs; β- and δ-COPs; γ -, ζ - and δ-COPs. Coatomers have a halflife of 28 h, although ζ -COP may exist in a stable form outside of the coatomer complex. COP-II coat components Sar1p Sar1p is also a small (24 kDa) GTP-binding protein. Its conversion into the GTP-bound form occurs through interaction with the integral membrane glycoprotein Sec12p [9], an event catalysed by Sec23p of the Sec23 complex [263]. Sec13/23 complexes Unlike the coatomer of COP-I vesicles there are two dimeric coat complexes which exist separately in the cytosol (see Table 3). The Sec13 complex (700 kDa) comprises Sec13p and Sec31p; both proteins contain numerous WD motifs [205]. The 400 kDa Sec23 complex consists of Sec23p and Sec24p [82]. The surface of COP-II vesicles has been visualized by deep-etch and rotary shadowing: unlike CCV it does not have a polygonal substructure, instead irregular clusters of 2 and 4 nm particles can be seen [211]. COP-vesicle formation and coat disassembly The recruitment of COP-I and -II coat proteins onto membranes appears to be similar. First, ARF1/Sar1p is attached, which does not require GTP hydrolysis. In the case of Sar1p, two proteins in addition to the GEF are involved: Sed4p and Sec16p. Then coatomers, or the Sec13/23 complexes, associate with the GTPbinding proteins. In the case of the coatomers this

occurs via β-COP [266]. Coatomers may also bind to the membrane directly via lecithin and/or phosphatidic acid, under the participation of phospholipase D [150], or through an interaction between a subset of coatomer subunits (α-, β 0 - and ε-COPs) and a dilysine (KKXX) motif (see below) in the cytoplasmic tail of certain transmembrane proteins [116]. A specific coatomer-binding transmembrane protein has recently been identified, p23, which has a short cytoplasmic domain containing a dilysine-type motif. Compared to Golgi membranes p23 is enriched 20-fold in COP-I vesicles [229]. The release of COP-I vesicles, in contrast to COPII vesicles, requires palmitoyl-CoA and ATP [175]. Another difference between COP-I and COP-II vesicles may exist; this deals with the timing of GTP hydrolysis and release of the coat proteins. Although ARF1 is concentrated in COP-I vesicles [52] and the prevention of coat protein dissociation by GTPγ S still allows their successful docking onto acceptor membranes (see above), Schekman and Orci [211] have proposed that in the case of COP-II vesicles Sar1p may even be released immediately upon completion of the coat assembly process. COP vesicle functions As far as COP-II vesicles are concerned, there appears to be general agreement that they function solely to transport in the anterograde direction from the ER to the Gapp. However, before they even fuse with the Gapp it would seem that they have the capacity to exchange their coat protein complexes for ARF and coatomers [203]. This occurs in a pre-Golgi or intermediate compartment [209]. What then happens to the COP-I vesicles is highly controversial, as can be read in the most recent editorial by Schekman and Mellman [212]. Evidence for COP-I vesicles operating in the retrograde direction The suggestion that COP-I vesicles are responsible for recycling of proteins from the Gapp back to the ER came originally from in vitro binding experiments [36, 125] and from an analysis of yeast COP mutants [116]. As mentioned above, it has been demonstrated that a subset of the COP-I coatomer can bind to an immobilized protein chimera containing a terminal dilysine motif. Moreover, evidence for the participation of δand ζ -COPs in the retrieval of ER proteins is now at hand [37]. It has also been shown that ret mutants,

61

Figure 3. In vitro system for the induction and isolation of COP-coated vesicles, as based on the procedures of Rothman, Schekman and coworkers. Step 1: incubation of donor membranes and cytosol (coatomers-Sec13/23 complexes) in the presence of an ATP-regenerating system and GTPγ S. Step 2: recruitment of coat proteins and vesicle budding. Step 3: removal of COP vesicles from target membranes through high-salt treatment. Step 4: separation of COP vesicles by isopycnic density gradient centrifugation.

which are incapable of retrieving membrane proteins with a terminating KKXX motif from the Gapp, were actually mutants of α-, δ-, and ε-COPs. An apparent exception to the rule seemed to be Emp47p which, although it possesses a dilysine motif in its cytoplasmic domain and continually recycles between the ER and Golgi in yeast, remains in the Golgi in ret1-1 (α-COP) mutants, which were otherwise defective for other dilysine proteins [219]. However, Emp47p is mislocalized to the vacuole in sec21-1 (γ -COP) mutants [117] suggesting that COP-I vesicles might mediate the retrieval of Emp47p from a more distal Golgi compartment. A similar retrograde transport function for COP-I vesicles in retrieving processing enzymes from maturing cisternae within the Golgi stack has also been proposed [62, 72]. Evidence for COP-I vesicles operating in the anterograde direction Three lines of evidence are normally given in support of an anterograde transport function for COP-I vesicles. Firstly, BFA, which prevents coatomer bind-

ing to ARF1 (see above), effectively blocks secretion in mammalian cells in vivo and leads to the fragmentation of the Gapp [123]. In the in vitro CHO Golgi transport system BFA does not prevent the transport of VSV-G protein [241]. However, Schekman and Mellman [212] have attempted to explain this contradictory observation in terms of a non-vesicular transport brought about by the BFA-induced formation of intercisternal tubular linkages. Secondly, yeast coatomer mutants show impaired secretion [92], although this effect is selective and is now interpreted in terms of a very tight coupling between COP-I and COP-II transport at the cis Golgi [61]. Thirdly, microinjection of β-COP antibodies inhibits anterograde secretory and membrane traffic [171]. Do COP-I vesicles facilitate transport in both anteroand retrograde directions? Orci et al. [161] have recently presented very impressive immunocytochemical data pertaining to this question. Using antisera directed against secretory (proinsulin), plasma membrane (VSV-G) proteins and the

62 KDEL receptor, as well as against β-, ε- and ζ -COPs, they investigated the relative distribution of anteroand retrograde cargo molecules in Golgi-associated COP-I vesicles in pancreatic endocrine cells in situ. In addition, they examined the contents of COP-I vesicles formed by isolated Golgi membranes. In each case two separate populations of COP-I vesicles were observed: one for the anterograde (proinsulin, VSV-G protein), and the other for the retrograde (KDEL receptor) traffic. Evidence for a co-localization of proinsulin and the KDEL receptor in the same COP-I vesicle was not obtained. KDEL receptor-containing COPI vesicles were found predominantly at the cis pole of the Golgi, whereas proinsulin-containing COPvesicles seemed to bud from every level of the Golgi stack. COP-vesicles in plants Although profiles of budding COP-like vesicles (60– 90 nm diameter, sometimes with a visible nap-type coat) on the cisternae of plant Golgi stacks are frequently to be seen in the published literature (see for example Figure 4, and [14]), actual evidence that they do represent COP vesicles is not yet available. Nor have any publications appeared relating to their isolation or in vitro induction from plant extracts. On the other hand, plant cells are known to respond to BFA treatment in a manner similar, albeit not identical, to mammalian cells (reviewed by [207]). Homologues for ARF1 [83, 189], Sar1p [41, 54], and Sec12p [54] have also been recorded. Thus, it is highly probable that COP vesicles, like CCV, are present in plants, and it is surely only a question of time before this fact is demonstrated. Our group has recently started investigations towards this goal, by generating antisera against GSTfusion proteins of Arabidopsis Sec21p (γ -COP), and Sec23p (COP-II coat complex) homologues. In comparison to antisera prepared against the corresponding yeast antigens (kindly provided by R. Schekman), our plant COP antibodies are much more effective in recognizing polypeptides of the expected correct molecular mass in cytosolic extracts from cauliflower inflorescence (see Figure 5). The opposite is also true when yeast and cytosolic extracts are probed with AtSec21p antibodies. AtSec23p antibodies, in contrast, recognize weakly the appropriate antigens in yeast and brain cytosol. We have already probed subcellular fractions from cauliflower inflorescence with these antisera and have obtained results which are conform

with the concept that COP-I vesicles are generated by the Gapp, while COP-II vesicles are formed at the ER (Movafeghi and Robinson, unpublished data). Dense vesicles Occurrence, morphology, and sites of formation There is growing evidence in support of the notion that plant cells may possess different types of vacuole in the same cell (see Neuhaus and Rogers, this volume). A logical consequence of this observation is the expectation that there should be more than one type of Golgi-derived vesicle involved in vacuolar protein transport, and this is indeed the case in seed tissues, which in addition to having lytic-type vacuoles also develop a second type of vacuole for the purpose of accumulating storage proteins (see Muentz, this volume). When aggregated, storage proteins are highly osmiophilic which makes them easy to detect in thin sections in the electron microscope. Electron opaque deposits of this type are not only seen in the protein storage vacuole, but are also present as the core of DV (Figure 6a) and have been described for the endosperm of cereal grains (e.g. wheat [102, 118]) and castor bean [71], as well as for the cotyledons of pumpkin [70] and various legumes (e.g. common bean [151], garden bean [12], pea [73, 88, 193] and soybean [78]). The DV in castor bean endosperm and in pumpkin cotyledons have a diameter of around 300 nm and appear to be generated at the ER and bypass the Gapp. Although Golgi marker enzymes were not measured this interpretation is supported by radiolabelling experiments in which precursor proteins were seen to rapidly chase out of the ER and into a high density, DV-containing fraction after only 30 min [57, 69]. In the other cases mentioned above the DV are clearly formed at, and bud from the Gapp. The best example investigated so far is that of the pea cotyledon DV [88, 191, 192]. These are quite uniform in diameter (130 nm; Figure 6), and when released from the Gapp appear to be smooth surfaced. DV begin to be formed at the cis-most Golgi cisternae, and, based on the staining intensity of their contents, seem to undergo some sort of maturation as they progress through the Golgi stack (Figure 6d; [192]; for mechanisms of intra-Golgi transport, see Faye, this volume). While still being attached to tubular elements at the TGN, most if not all DV are partially capped with a clathrin coat from which a CCV is later formed (Figure 6b, c). This is structurally an analogous situation

63

Figure 4. Putative COP-coated vesicles in plants. a, b. Golgi apparatus in Chlamydomonas reinhardtii with associated vesicles. In a, (COP-II ?) vesicles can be seen budding at the endoplasmic reticulum, which lies immediately opposite the cis (c) pole of the Golgi stack. In b, the plane of section is parallel to the major axis of the cisternae, thereby revealing numerous (COP-I ?) vesicles at the periphery. c. Golgi apparatus in developing pea cotyledons showing a dense vesicle (arrowhead) at the trans pole, and possible COP-vesicles (arrows). Bars = 200 nm.

to that recorded for the immature secretory granules of neuroendocrine cells and cells of exo-and endocrine glands in mammals [47, 112]. Preliminary characterization DV have been successfully isolated from developing castor bean endosperm and pumpkin and pea cotyle-

dons, and, despite their different sizes (see above), have a common isopycnic density in sucrose of 1.22– 1.24 g/ml [30, 57, 69, 87]. They have a mixed cargo: in pea DV both types of storage globulins (vicilin and legumin) are present in their unprocessed, proform in the same vesicle [87, 88]. In the case of pumpkin and castor bean DV, the storage globulins are transported

64

Figure 5. Cross reactivities of polyclonal Sec21p and Sec23p antisera prepared from yeast and Arabidopsis antigens. Each antiserum was tested by western blotting on cytosolic extracts prepared from porcine brain (B), cauliflower inflorescence (C), and yeast (Y). Details of the cDNA clones, and the fusion proteins used for the antibody production are given below.

together with their specific, maturation proteases [70, 71]. The latter, in the main cysteine endoproteases [142], are in an inactive form in the DV, and become active via a pH-dependent autocatalytic process in prevacuolar or vacuolar compartments. Unlike CCV, which are enriched in the vacuolar sorting receptor BP-80 (see above), DV are without BP-80 but carry the typical protein storage vacuole aquaporin, α-TIP (Figure 7; [87, 193]). Mechanism of storage protein segregation and aggregation In contrast to lysosomal and vacuolar acid hydrolases which are sorted from other elements of the secretory pathway at the TGN, the sorting machinery for storage globulins already operates in the cis-cisternae of the Gapp. Based on the observation that complex glycoproteins, but not vicilin and legumin, are detectable with the immunogold method in the central part of the cisterna [88], the separation of newly arrived proproteins from the rest of the lumenal contents and their transport to the periphery of the cisternae, where they aggregate, must be a highly efficient process. Whether this aggregation event is chaperonemediated, or whether it occurs spontaneously after a critical concentration is exceeded remains to be elucidated. It is, however, different to the situation in vitro [46] and in transgenic plants where it has been shown that prolegumin cannot form oligomers larger than trimers unless it is processed into the mature form, which occurs in the vacuoles [97]. By com-

parison, prolegumin present in hexamers has been demonstrated in protein storage vacuoles isolated from developing pea cotyledons [86], and, as already mentioned, prolegumin in pea cotyledon DV is exclusively present in its unprocessed form. Considering the highly aggregated state of the proteins in the DV it is also uncertain whether a ‘classical’ type of ligand/receptor interaction with a 1:1 stoichiometry is responsible for the initial sorting event, despite the fact that prolegumin appears to possess a vacuolar sorting signal which is capable of directing reporter constructs into the vacuoles of tobacco leaves and seeds [204]. Aggregation-mediated protein sorting as a means to explain the formation of immature secretory granules in various animal cells (recently reviewed by [243]) may, however, also apply to the DV of plant cells (see Figure 8). According to this scenario, secretory proteins (here the aggregated content of the immature secretory granules) interact with a hydrophobic subpopulation of the same proteins which are already tightly attached to the membrane (in this case the TGN). These proteins are thought to act as a nucleus triggering the aggregation of the regulated secretory proteins, thereby inducing the budding of the immature secretory granule [177]. Both membrane association and aggregation are possibly the result of hydrophobic protein-protein interactions. The existence of a subpopulation of membrane-associated secretory proteins points to the presence of a new type of sorting receptor, which, in contrast to the mannose 6-phosphate receptor or the yeast Ypt10p need not

65

Figure 6. Dense vesicles (DV) in developing pea cotyledons. a. Overview of a Golgi-rich area. Osmiophilic DV are indicated with arrowheads, clathrin-coated vesicles (CCV) with arrows. b, c. Sectional profiles of CCV budding from DV. d. DV mature across the Golgi stack: those attached to the cis (c) cisternae have less osmiophilic contents than those at the trans pole. e. Isolated DV fraction. Bars = 500 nm (a), 100 nm (b-e).

66

Figure 7. Characterization of dense vesicles (DV) by immunocytochemistry on cryo-sections. a. Positive labelling of DV (arrowheads) with α-TIP antibodies. b. Negative labelling with BP-80 antibodies, which positively label the cisternae. Bars = 200 nm.

be recycled to the TGN in order to maintain correct sorting for a longer period of time. In keeping with this hypothesis, pea prolegumin is much more hydrophobic than mature legumin and is also tightly bound to membranes of the secretory pathway. This association persists even after solubilization of most of the remaining proteins with digitonin [86]. The same behaviour is also shown by vicilin, the other major storage protein in pea seeds (Hinz, unpublished results). By contrast, mature legumin is only loosely associated with the membrane of isolated protein bodies [86]. A second feature of DV indirectly supportive of this hypothesis is the formation and budding of CCV. In immature secretory granules, but not in mature secretory granules lysosomal hydrolases are still present, and AP-1 have been detected at the membrane surface [47, 112] indicating that this second vesiculation event serves to retrieve missorted acid hydrolases via the mannose 6-phosphate receptor.

How vesicles recognize their fusion target: the SNARE hypothesis The segregation and collection of cargo molecules and their packaging into specific transport vesicles only makes sense when the vesicle can find its correct target. Five years ago an hypothesis was put forward to explain just how this feat of intracellular navigation

might be accomplished: this is the SNARE concept [228]. In the meantime, a large body of evidence, obtained mainly on mammalian and yeast cells, but also on plants, has accrued in support of this hypothesis, and several excellent reviews specifically devoted to this subject have been published [e.g. 19, 170, 176]. Unfortunately, and especially for the plant scientist who is unfamiliar with this field, there is no uniform nomenclature in use: the mammalian and yeast researchers each have their own jargon. The SNARE concept finds its origin in the classic experiments on in vitro protein transport performed by Rothman and others in the 1980s. These early studies established that two soluble factors, NSF (Nethylmaleimide-sensitive factor) and SNAP (soluble NSF attachment protein), were absolutely necessary for successful vesicular transport (i.e. resulting in fusion). A subsequent search for the proteins with which NSF and SNAP interact led to the discovery of two types of membrane protein: one set characteristic for the vesicle and termed v-SNARE (vesicle SNAP receptor), the other called t-SNARE (target SNAP receptor). Other players now recognized as belonging to the team are the SNARE blockers, the Rab GTPases, and the Rab effectors. A possible way in which they work together to facilitate vesicle docking and fusion is presented in Figure 2F.

67 SNAPs Three SNAP isoforms (α, β and γ ), each with a molecular mass around 37 kDa, have been identified in mammalian cells [32] and shown to bind to Golgi membranes in vitro [257]. This binding requires the presence of assembled SNARE complexes [228] and may involve other proteins as well [134]. The gene for α-SNAP in yeast is SEC17 [67]. To our knowledge, SNAP homologues have not yet been identified in plants. SNAREs

Figure 8. Possible mechanism of protein sorting and vesicle formation, as based on the model of Thiele et al. [243], but modified for the specific case of the Golgi apparatus in pea cotyledons. Segregation of storage proteins is the consequence of a ‘domino’ effect started by a hydrophobic subpopulation of membrane-binding storage proproteins. TGN, trans Golgi cisterna/network; IDV, immature dense vesicle (containing storage proprotein and missorted acid hydrolases); DV, dense vesicle (containing storage proprotein); CCV, clathrin coated vesicle containing acid hydrolases); SV, secretory vesicle; , secretory proteins; , vacuolar acid hydrolases; , storage proproteins; , membrane-attached subpopulation of storage proproteins.

#





NSF NSF is a trimeric ATPase with subunits of around 100 kDa, which exists in membrane-bound and cytosolic forms [258]. It is a member of a large class of ATPases which catalyze protein-protein interactions [136]. In mammalian cells a hexameric NSF variant, p97, has been identified [172] and shown to be required for the reassembly of mitotic [184] or inhibitor [2] dispersed Golgi membranes. The gene for NSF in yeast is SEC18 [98], and mutants of this gene reveal numerous interruptions in various intracellular transport pathways [66]. An homologue in yeast, Cdc48p, is absolutely required for homotypic ER fusion [113]. Interestingly, an increase in v-SNARE complexes has been detected in sec18 mutants [120] lending credence to the idea that NSF is involved in the dissociation, rather than the assembly of the v- and t-SNARE complex. On the other hand there is evidence that, at least for homotypic membrane fusions, NSF might also function in a predocking stage [149]. As far as we are aware only one NSF homologue has been reported in plants [95], and this is a plastid translocation factor.

SNAREs were originally found by eluting immobilized complexes prepared from NSF, α- and γ -SNAPs, plus detergent-solubilized bovine brain membrane proteins with Mg2+ -ATP [228]. ATP hydrolysis results in the release of NSF and the SNAPs, leaving behind a 7S complex which contains three synaptic proteins: synaptobrevin (also known as VAMP), a component of the synaptic vesicle membrane and hence a v-SNARE; syntaxin and SNAP-25 (synaptosomeassociated protein of 25 kDa, not to be confused with α-SNAP), both components of the presynaptic PM, and hence t-SNAREs. It turns out that synaptobrevin and syntaxin are members of gene families with representatives not only in mammalian cells [20], but in yeast [56] and plants [11, 127] as well. Common to nearly all SNAREs is that they have highly conserved and hydrophobic domains at their C-terminus, the latter anchoring them in the membrane. Pelham [170] has pointed out that the existence of eight syntaxin homologues can be predicted from the yeast genome sequence, six of which have been identified: Ufe1p (possibly Sec20p as well) is an ER t-SNARE, Sed5p and Tlg1/2p are t-SNAREs of early and late Golgi compartments, PEP12p is an endosomal/prevacuolar t-SNARE, Vam3p a t-SNARE of the vacuole, and Sso1/2p are PM t-SNAREs. A number of the v-SNAREs involved in ER to Golgi traffic have been identified: four v-SNARES (Bet1p, Bos1p, Sec22p, Ykt6p) are known to complex with Sed5p [230]. Interestingly, Vti1p a Golgi v-SNARE interacts with both PEP12p and Sed5p (von Mollard, personal communication), which is indicative of recycling from the TGN, in addition to its function in separating secretory from vacuole-destined products (see also [170]). Three syntaxin homologues in plants have been identified so far. One is a PEP12p homologue in Arabidopsis which can functionally complement for

68 pep12 in yeast mutants [11]. This t-SNARE has been localized to an undefined, post-Golgi structure in Arabidopsis [35]. Another one is the KNOLLE protein, also from Arabidopsis [127]. Mutations in the KN gene have serious consequences in cytokinesis and cell wall formation. This defect appears to be caused by the inability of Gapp-derived vesicles to fuse with one another during cell plate formation. This could, therefore, be an example of homotypic membrane fusion for which the KNOLLE protein is the necessary v/t-SNARE. The third example is AtVam3p which is implicated in vacuole biogenesis in Arabidopsis [208]. Two Arabidopsis homologues to the v-SNARE Vti1p (with 30% identicity to yeast Vti1p) have also been recently identified (von Mollard and Raikhel, personal communication). SNARE blockers If v-and t-SNARE interactions are not somehow regulated, there is a danger of uncontrollable and massive membrane fusions. To avoid this the SNAREs appear to be in a latent form and only become activated during vesicle formation. While coat proteins may affect SNARE accessibility some other proteins have been described which seem to bind to SNAREs. Thus, Sec1p in yeast [1] and its mammalian equivalent, munc18p [74], seem to protect PM t-SNAREs, and Sly1p blocks Sed5p [230]. A v-SNARE interacting protein from neuronal extracts, synaptophysin [53], has also been described. Whether these blocking proteins are completely dissociated from the SNARE complex after v-and t-SNARE interaction is unclear, but their existence means that SNARE recognition and interaction are more complicated than they seem (see [176] for a discussion). Rabs The correct pairing of two SNARE proteins is mediated by the action of Rab (Ypt) GTPases. Although Rabs do not seem to be a constituent of SNARE complexes, there is evidence that they are required for v- and t-SNARE interactions [230]. Thus, it is now known that Ypt1p transiently interacts with the t-SNARE Sed5p, thereby displacing the blocker Sly1p [128]. Up to now 44 members of the Rab family have been described for mammalian cells, associated with defined compartments of the secretory and endocytic pathways, and 11 have been identified in yeast (for recent reviews see [153, 233]). On the plant side, numerous Rab homologues have been reported (i.e. [15,

83, 94]). Alone 29 members of this protein family homologous to Rab1, Rab2, Rab5, Rab7, Rab8 and Rab11 have been recently identified in developing root nodules of Lotus japonicus [24]. Rabs are monomeric proteins of about 25–30 kDa and possess four conserved sequence motifs named G1, G3, G4 and G5. Located between the G1 and G3 domains is the G2 or so-called effector region, which is unique to each subclass, and which is important for the specificity of the downstream interactions of the subclasses. A cysteine-containing isoprenylation site is additionally located at the C-terminus [154]. Isoprenylation, which is necessary for membrane anchoring, is catalysed by the enzyme Rab geranylgeranyl transferase (GGTase). This reaction is further supported by the Rab escort protein (REP) [153], which binds and presents the unprenylated or monoprenylated precursor to the GGTase. During budding and fusion of transport vesicles Rab cycles between the donor and acceptor membranes. This so-called Rab cycle [153] begins in the cytosol with Rab (in the GDP-bound form) complexed with the Rab GDP-dissociation inhibitor (GDI). The latter prevents indiscriminate membrane binding of the Rab-GDP form due to exposed lipids. Upon reaching the donor membrane the Rab-GDP/GDI complex is dissociated through the action of GDI-displacement factors, allowing free Rab-GDP to bind to the membrane. Immediately after membrane attachment the bound GDP is exchanged by GTP, a reaction catalysed by the GEF. This renders the bound Rab-GTP inaccessible to the action of the GDI, which otherwise would again interact with the bound Rab-GDP. After docking of the vesicles to the acceptor membranes the GTP is hydrolysed with the help of the GTPase activating protein (GAP). Now the GDI can remove the Rab-GDP from the acceptor membrane allowing the cycle to start again. Since Rab-GGTase [126], and homologues for GDI [21, 247, 264] and GAP-like activity [8] have been identified in plants it is a safe assumption that the Rab cycle functions in the same way as it does in mammalian and yeast cells, and that vesicular transport in plants is also under the control of Rab proteins. Rab effectors As pointed out in recent reviews [153, 176, 233], there is now evidence for the participation of extra Rab-binding proteins, such as Rabex-5, Rabaptin 5, Rabphilin, Rim, Uso1p(p115) in vesicle docking [45, 91, 225, 237, 254]. These Rab effector molecules,

69 which appear to have long stretches of α-helices, bind to the variable G2 domain of Rab proteins. It is therefore conceivable that through coiled-coil interactions between opposing Rab effectors vesicles may be ‘caught’ by target membranes thereby allowing the SNAREs to interact.

12.

13.

14.

Acknowledgements

15.

Work from our group reported in this review was supported by the Deutsche Forschungsgemeinschaft (SFB 523; Teilprojekt A7). We thank Stephan Hillmer for preparing the cryosections, and Peter Pimpl and Bernd Raufeisen for the drawings.

16. 17.

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