Formation and maintenance of the Golgi apparatus in plant cells.

CHAPTER SIX

Formation and Maintenance of the Golgi Apparatus in Plant Cells Yoko Ito*, Tomohiro Uemura*, Akihiko Nakano*,†,1

*Department of Biological Sciences, Graduate School of Science, University of Tokyo, Bunkyo-ku, Tokyo, Japan † Live Cell Molecular Imaging Research Team, RIKEN Center for Advanced Photonics, Wako, Saitama, Japan 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Features of Plant Golgi as Compared to Animal and Yeast Cells 2.1 Morphology 2.2 Dynamics 2.3 Functions 3. ER–Golgi Transport 3.1 COPI vesicle 3.2 COPII vesicle and ERES 3.3 ERGIC 4. Intra-Golgi Trafficking 4.1 Models of intra-Golgi trafficking 4.2 Tethering and fusion of intra-Golgi COPI vesicles 5. Formation and Maintenance of Stacked Structure 5.1 Golgi apparatus as stacked cisternae 5.2 Search for the “Golgi stacking factors” 5.3 Golgi biogenesis upon mitosis 5.4 Golgi regeneration after BFA treatment 6. Relationship Between the Golgi Apparatus and the TGN 7. Concluding Remarks Acknowledgments References

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Abstract The Golgi apparatus plays essential roles in intracellular trafficking, protein and lipid modification, and polysaccharide synthesis in eukaryotic cells. It is well known for its unique stacked structure, which is conserved among most eukaryotes. However, the mechanisms of biogenesis and maintenance of the structure, which are deeply related to ER–Golgi and intra-Golgi transport systems, have long been mysterious. Now having

International Review of Cell and Molecular Biology, Volume 310 ISSN 1937-6448 http://dx.doi.org/10.1016/B978-0-12-800180-6.00006-2

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extremely powerful microscopic technologies developed for live-cell imaging, the plant Golgi apparatus provides an ideal system to resolve the question. The plant Golgi apparatus has unique features that are not conserved in other kingdoms, which will also give new insights into the Golgi functions in plant life. In this review, we will summarize the features of the plant Golgi apparatus and transport mechanisms around it, with a focus on recent advances in Golgi biogenesis by live imaging of plants cells.

1. INTRODUCTION The Golgi apparatus is a single-membrane-bounded organelle in eukaryotic cells, and its pivotal roles in intracellular membrane trafficking are well established since its first description. The Golgi apparatus consists of several flattened membrane sacs called cisternae, which are stacked in layers like pancakes. In each Golgi stack, cisternae are polarized between the cis side, receiving cargo from the endoplasmic reticulum (ER), and the trans side, sending cargo forward to post-Golgi organelles (Mellman and Warren, 2000). Along this axis, many glycosylation enzymes are arranged as a gradient, so that cargo proteins can be modified sequentially by passing across the Golgi from cis to trans (Dunphy and Rothman, 1985; Moore et al., 1991). This stacked structure, which makes the Golgi apparatus “one of the most easily identified components of the cytoplasm” (Porter, 1961), is the biggest and unique morphological feature of this organelle. Although it has been attracting many biologists for a long time, little is known about how this structure is formed and maintained. In this review, we summarize the knowledge about the plant Golgi morphology, dynamics, functions, and underlying molecular mechanisms in comparison with yeast and animals. In addition, we highlight the recent data by live imaging approaches to reveal the biogenesis process of the stacked structure and its relationship with the ER–Golgi trafficking.

2. FEATURES OF PLANT GOLGI AS COMPARED TO ANIMAL AND YEAST CELLS 2.1. Morphology When Camillo Golgi first reported about the peculiar structure he had observed in the Purkinje cells of the cerebellum, he designated the structure as “apparato reticolare interno (internal reticular apparatus)” with fine and

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elegant network (Golgi, 1898). He also mentioned in his later writing that the reticular apparatus occupied the perinuclear zones and the nucleus was somehow embraced by it (Bentivoglio, 1998). However, these morphological features of the “classical Golgi” were soon questioned by the researchers who tried to identify the Golgi apparatus in invertebrate cells. When they applied Golgi’s staining method to invertebrate cells like insects or molluscs, discrete structures (granules, curved rods, and filaments) were observed (Beams and King, 1932). At that time, because of the absence of electron microscopic techniques, the existence of the Golgi apparatus itself was still under intense debate even in mammalian cells (Beams and Kessel, 1968; Bentivoglio, 1998). Moreover, since the appearance of the invertebrate Golgi apparatus was quite different from that of mammalian nerve cells, their homology drew much controversy (Beams and King, 1932). In the midst of the debates, it was suggested to call the discrete Golgi materials in insect cells as “dictyosomes,” the term that had been originally used to describe the pieces into which the Golgi apparatus fragmented during mitosis in vertebrate cells (Bowen, 1920). As regards the Golgi apparatus in plants, among suggestions that put the “vacuolar system” or the plastids as the homologs of the Golgi (Beams and Kessel, 1968), Bowen found that there were intracellular structures resembling the dictyosomes in many kinds of plant cells. They were independent from all the cytological structures previously described and he called them “osmiophilic platelets” (Bowen, 1928). In addition to the similarity in their appearance, it was demonstrated that the “osmiophilic platelets” occupied the same relative position as did the Golgi apparatus in animal cells upon ultracentrifugation (Beams and King, 1935a,b). In 1951, the first application of electron microscopy to the Golgi apparatus finally confirmed its existence (Dalton, 1951). Soon after, with the development of techniques for electron microscopic observation, the fine structure of the Golgi apparatus in vertebrate cells was revealed: concentrically arranged numerous flattened sacs of smooth-surfaced membranes with small vesicles (Dalton and Felix, 1954; Sjo¨strand and Hanzon, 1954). This characteristic structure of the Golgi apparatus was also found in insect cells (Beams et al., 1956), and of course, in various kinds of plant cells (Beams and Kessel, 1968). After all the half-century of discussion, the Golgi apparatus became to be recognized as an intracellular structure, which is conserved among most eukaryotic cells. Today, the term “osmiophilic platelets” is no longer in use, however, “dictyosome” still remains as a word to represent the Golgi stacks especially in plant sciences.

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As we can see from the history of the Golgi discovery, in spite of the conservation of their fine stacked structure at the electron microscopic level, the morphology of the whole Golgi apparatus is quite different among species. The plant Golgi apparatus consists of numerous, sometimes many hundreds of individual stacks of disk-like cisternae, and they are usually dispersed throughout the cytoplasm (Fig. 6.1A; Dupree and Sherrier, 1998; Hawes et al., 2008b). Each unit has a clearly stacked structure and cis–trans polarity, which can be confirmed either by the fine architecture under the electron microscope or by the localization of the marker proteins such as glycosylation enzymes and proteins involved in the trafficking machinery (Dupree and Sherrier, 1998; Ito et al., 2012; Schoberer and Strasser, 2011; Staehelin and Kang, 2008). This plant-style Golgi organization is also observed in protozoa, fungi (such as Schizosaccharomyces pombe and Pichia pastoris), and invertebrates (Wei and Seemann, 2010). On the other hand in vertebrate cells, as Camillo Golgi observed, the Golgi apparatus typically exhibits a twisted network, which is called the “Golgi ribbon” after its appearance under the light microscope

Figure 6.1 Comparison of the Golgi organization among eukaryotes. (A) In plant cells, individual Golgi stacks with clearly stacked structure are spread in the cytoplasm. Actin filaments are often arranged along the ER, and the Golgi stacks move on them by myosin motors. (B) In mammalian cells, microtubules are organized in a radial pattern around the centrosome. The Golgi stacks are centralized by dynein motors on the microtubules and interconnected to form the Golgi ribbon. ERGIC is formed near the ER, and ERGIC itself or transport vesicles from it travel to the Golgi ribbon along the microtubules. (C) In S. cerevisiae, the Golgi apparatus rarely forms stacks, and individual cis, medial, and trans cisternae scatter in the cytoplasm.


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(Fig. 6.1B; Lowe, 2011; Wei and Seemann, 2010). The Golgi ribbon is closely associated with the centrosome (or the microtubule-organizing center; MTOC), which is usually located near the nucleus in nonpolarized mammalian cells (Rios and Bornens, 2003). Because the Golgi ribbon is fragmented into many pieces with typical stacked organization (the ministacks) by depolymerizing microtubules (Polishchuk et al., 1999), it can be thought that the Golgi ribbon is an interconnected assemblage of the ministacks, which are very similar to the “dictyosome”-style Golgi in nonvertebrate organisms. Moreover, even in mammals, the Golgi apparatus naturally exhibits isolated stacks in certain types of cells (such as oocytes and skeletal muscle cells; Matteis et al., 2008). Hence, each discrete stacked structure as a minimum unit of the Golgi is thought to be fundamentally conserved among eukaryotes. However, there are exceptions. In the budding yeast Saccharomyces cerevisiae, the Golgi membranes rarely form stacks and scatter in the cytoplasm as individual cisternae, which exhibit tubular network-like structure. Each cisterna can be defined as cis, medial, and trans by the localization of the marker proteins (Fig. 6.1C; Matsuura-Tokita et al., 2006; Preuss et al., 1992; Rambourg et al., 1993). Moreover, in certain parasitic protists, the stacked Golgi structure was never observed and thus these species were considered as “Golgi-lacking” organisms ( Je´kely, 2008; Mowbrey and Dacks, 2009). It is now thought that species without Golgi stacks such as S. cerevisiae have evolved from the ancestors with stacked Golgi according to phylogenetic analyses (see Section 5.1; Mowbrey and Dacks, 2009). Although major progress has been made in the Golgi research by using mammalian cells, the complexity of the Golgi ribbon makes it difficult to observe the stacked structure in living cells by light microscopy. S. cerevisiae greatly contributed to resolve the molecular mechanisms of intracellular trafficking as a model organism (Schekman, 1992), and moreover, Ben Glick’s group and ours took advantage of the unstacked feature of the Saccharomyces Golgi to demonstrate the cisternal maturation (see Section 4.1; Losev et al., 2006; Matsuura-Tokita et al., 2006). However, we cannot investigate cisternal stacking in S. cerevisiae. By contrast, thanks to the simple and elegant structure, the Golgi stacking can be easily studied by light microscopy in plant cells. By introducing multiple marker proteins, each of which has distinctive intra-Golgi localization and a different colored fluorescent tag, the plant Golgi stacks are visualized as many punctate structures with the markers localized in layers (Fig. 6.2). On top of this, even though the size of the plant Golgi stacks can vary between species, cell types,

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Figure 6.2 Visualization of the plant Golgi stacks in living cells. (A) Confocal image of a tobacco BY-2 cell expressing GFP-AtSYP31 (cis-Golgi, green) and ST-mRFP (trans-Golgi, magenta). Arrowheads indicate the Golgi stack used for the analysis in B. Scale bar, 10 mm. (B) The fluorescence profile along the arrow across a Golgi stack.

and cell conditions, they are generally larger than the counterparts in other multicellular model organisms with dispersed Golgi [in diameter under electron microscopes, 370 nm in tissue culture S2 cells of Drosophila melanogaster (Kondylis and Rabouille, 2009), 200 nm in body-wall muscles and 500–800 nm in macrophage-like coelomocytes in Caenorhabditis elegans (Luo et al., 2011), 800 nm in tobacco BY-2 cultured cells (Nebenfu¨hr et al., 1999)]. The morphological features of the plant Golgi provide a great advantage for detailed analysis of the stacked structure by live imaging.


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2.2. Dynamics In plant cells, the majority of organelle movement is actomyosin dependent (Sparkes, 2011). Owing to the development of live imaging techniques, it has become clear that the Golgi apparatus is not an exception; the Golgi movement is greatly inhibited by actin disrupting drugs such as cytochalasin D and Latrunculin A, and by the myosin ATPase inhibitor 2,3-butanedione monoxime (Boevink et al., 1998; Nebenfu¨hr et al., 1999). In addition, by confocal microscopy, movement of the Golgi stacks can be observed along the actin filaments in living cells (Fig. 6.3). From the observation in tobacco BY-2 cells whose Golgi stacks are visualized by cis- and trans-Golgi markers tagged by different colors, the stacks are suggested to have a slight preference to direct the cis side ahead during the movement along the actin filaments, although the trans side can also

Figure 6.3 Plant Golgi stacks move along actin filaments. Time-lapse observation of a tobacco BY-2 cell expressing GFP-AtSYP31 (cis-Golgi, green), ST-mRFP (trans-Golgi, red), and Lifeact-Venus (actin filaments, blue). The Golgi stack indicated by the arrowheads moves toward the right. The indicated times mean the elapsed time. Scale bar, 5 mm.

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go forward and the stacks sometimes roll to change the leading side (Madison and Nebenfu¨hr, 2011). Microtubules, the other major cytoskeletal system, is not required for long-range movement of the plant Golgi stacks, because the treatment by microtubule disrupting drugs such as nocodazole, colchicine, and propyzamide does not have significant effects (Nebenfu¨hr et al., 1999). Plant myosins are classified into classes VIII and XI; the latter is structurally similar to class V myosins of metazoa and fungi, and is thought to share the common function in organelle movement (Avisar et al., 2008; Sparkes, 2011; Tominaga and Nakano, 2012). Several class XI myosins are shown to have overlapping functions and contribute to the Golgi movement to various degrees from knockout analyses, RNAi, and dominant negative inhibition in Arabidopsis and tobacco cells (Tominaga and Nakano, 2012). Observation of the Golgi movement in Arabidopsis knockout lines has recently suggested that myosin XI-K is the main contributor among them (Avisar et al., 2012). The dynamic flow of the cytoplasm including many organelles in plant cells has been long known as the cytoplasmic streaming (Shimmen and Yokota, 2004). Recent work has suggested that the unidirectional stream of the ER meshwork, which is driven primarily by myosin XI-K, hauls the cytosol and thus causes the cytoplasmic streaming (Ueda et al., 2010). The plant Golgi stacks may participate in this passive cytoplasmic streaming as well; however, their movement is not simple. The Golgi stacks exhibit stop-and-go motions; they alternate between pause for seconds to minutes and rapid directed movement, for example, with the velocity up to 7 mm/s in highly vacuolated Arabidopsis root cells (Akkerman et al., 2011; Boevink et al., 1998; Nebenfu¨hr et al., 1999). It is suggested that this Golgi motility pattern depends on the actin configuration (Akkerman et al., 2011), and that cortical microtubules establish “landmark sites,” which cause frequent pause of organelles not only the Golgi apparatus but also RNA processing bodies, peroxisomes, mitochondria, and the tubular ER junctions (Hamada et al., 2012), but the functional significance of such features of cytoskeletons is not clear. The Golgi motions were initially related to ER–Golgi transport; the Golgi stacks were proposed to receive cargo from the ERES (ER exit/export site) during stationary phases, because the Golgi stacks traveling in the same track appeared to stop occasionally at the same positions (Nebenfu¨hr and Staehelin, 2001; Nebenfu¨hr et al., 1999). This model was supported by the result that the inhibition of ER–Golgi transport


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by overexpression of a dominant negative form of RABD2A reduced the Golgi movement (Batoko et al., 2000). However, evidence to argue against this model was reported by fluorescence recovery after photobleaching (FRAP) analysis, which showed that ER-to-Golgi protein transport occurred while the Golgi stacks were moving (daSilva et al., 2004). This question goes together with the debate about the relationship between the Golgi stacks and ERES in plant cells (see Section 3.2.4), and is still under controversy. In addition, cellulose synthase complexes are suggested to be inserted to the plasma membrane from the Golgi, when the Golgi stacks pause on cortical microtubules (Crowell et al., 2009; Gutierrez et al., 2009). More experimental data are awaited, which visualize the Golgi stacks and their cargo transport. In contrast to plant Golgi, which uses actin filaments as their highways (Figs. 6.1A and 6.3), mammalian Golgi uses microtubules for long-range transport, although actin filaments also take parts in the Golgi structural organization and carrier vesicle formation (Egea et al., 2013). Mammalian cells have radial organization of microtubules around the centrosome with minus-ends anchored at the centrosome and plus-ends extending to the cell periphery (Fig. 6.1B; Egea and Rıós, 2008). The minus-end-directed motor proteins, mainly dynein-1, collect the Golgi stacks along the microtubules and form the Golgi ribbon (Brownhill et al., 2009). A common feature of the cell types that do not have a continuous Golgi ribbon (from plant cells to mammalian oocytes and myotubes) is the absence of the radial patterned microtubule around the MTOC, indicating that the microtubule organization plays important roles in Golgi ribbon formation (Matteis et al., 2008). In addition, mammalian Golgi itself has been reported to act as an MTOC to nucleate microtubules both in vitro and in vivo, and the Golgi stacks can selforganize into a single continuous unit using self-derived microtubules in cytoplasts without centrosomes (Su¨tterlin and Colanzi, 2010; Wei and Seemann, 2009). Mammalian Golgi and the centrosome–microtubule system functionally interact with each other, not only in interphase but also during mitosis, and such relationship is not found in organisms with dispersed Golgi stacks (Su¨tterlin and Colanzi, 2010). Mammalian Golgi ribbon is relatively stable, but both the Golgi and the centrosome reorient to the traveling direction in many cell types upon migration (Brownhill et al., 2009). Moreover, dispersion or mislocalization of the Golgi ribbon without affecting centrosome-derived microtubule organization perturbs directional cell migration, while normal

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secretion ability is not affected (Hurtado et al., 2011; Yadav et al., 2009). The reason why mammalian Golgi apparatus forms a ribbon and behaves as a single organelle is still unclear; however, from the facts above, it is widely thought that Golgi ribbon formation maximizes polarized secretion efficiency. Then, are the cells without Golgi ribbon unable to carry out polarized secretion? Of course they can, in alternative ways. In S. cerevisiae, the Golgi cisternae cluster just below the growing bud during the initiation of new daughter cells, and they are also concentrated near the site of cell wall synthesis during cytokinesis (Preuss et al., 1992). This Golgi clustering can be interpreted as transient Golgi ribbon formation near the secretion target, while the cisternae are not interconnected. In D. melanogaster, the dispersed Golgi stacks are almost always found in close association with ERES (Kondylis and Rabouille, 2009), and a thousand of the ERES–Golgi units are evenly distributed in oocytes (Herpers and Rabouille, 2004). Among them, only the ERES–Golgi units in the dorsal/anterior corner transport Gurken protein, whose luminal fragment is secreted locally at the dorsal/anterior corner to specify the anteroposterior and dorsoventral axes, whereas all the ERES–Golgi units in the cell are equal in transport ability. This polarized transport is suggested to depend on polarized localization of gurken mRNA combined with its local translation (Herpers and Rabouille, 2004). This indicates that the dispersed Golgi stacks could functionally differentiate to transport different proteins. In support of this, the subset of Golgi stacks are proposed to possess different glycosylation enzymes in a single Drosophila imaginal disk cell (Yano et al., 2005). Also in plant cells, the Golgi stacks are shown to localize in the region several microns behind the growing tips of root hairs and pollen tubes, while the extreme tips are occupied by vesicles including Golgi-derived secretory vesicles (Campanoni and Blatt, 2007; Dupree and Sherrier, 1998). This Golgi behavior might be similar to the Golgi clustering in S. cerevisiae for polarized secretion, but the association of the Golgi stacks with growing regions is not absolute in plant cells (Dupree and Sherrier, 1998). It is unclear whether the Golgi stacks are actively gathered, or their localization is just the consequence of the concentration of the cytoplasm and only the vesicles are targeted to the growing tips. Another major event that requires polarized secretion in plant cells is cell plate formation upon cytokinesis. It is well known that the Golgi–TGNderived vesicles carrying cell wall components are targeted to the division


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plane, and the cell plate is formed by fusion of these vesicles ( Ju¨rgens, 2004; Staehelin and Hepler, 1996). In the past without live imaging techniques, the plant Golgi stacks were thought to have no particular organization during mitosis (Hepler and Wolniak, 1984). However, live imaging observation in tobacco BY-2 cells revealed that the general Golgi streaming stops when the cells enter mitosis, and a subset of stacks begin to redistribute around the perinuclear region. Later in metaphase, they aggregate around the mitotic spindle and in the equatorial region under the plasma membrane. This latter region accurately predicts the future site of cell division, and has been named the “Golgi belt.” During cytokinesis, many Golgi stacks localize around the phragmoplast, an array of microtubules required for proper targeting of the vesicles for the cell plate (Nebenfu¨hr et al., 2000). Such accumulation of the Golgi stacks has been also observed in Arabidopsis shoot apical meristem cells, in which up to 80% of the stacks localize in close proximity of the phragmoplast in early telophase (Seguı´-Simarro and Staehelin, 2006). This transition of the Golgi localization is thought to reflect the centralization of the secretory pathway for rapid cell plate synthesis. Although the orientation of the Golgi stacks and their associated TGN toward the target presumably affects polarized secretion efficiency, any biased orientation of the Golgi– TGN units has not been observed (Seguı´-Simarro and Staehelin, 2006). It is still unknown whether the plant Golgi stacks functionally differentiate in a single cell, like the case of ERES–Golgi units in Drosophila cells.

2.3. Functions 2.3.1 Subcompartmentalization of glycosylation enzymes The Golgi apparatus plays central roles not only in cargo transport and sorting but also acts as an important biosynthetic compartment where the complex oligosaccharides moieties of glycoproteins and glycolipids are assembled. This Golgi function as the biochemical “factory” is fundamentally conserved among eukaryotes, albeit the actual modification steps may differ between species or cell types (Gomord et al., 2010; Song et al., 2011). A widely accepted view on the sugar chain modification process is the assembly-line model, in which sequentially ordered glycosylation enzymes elongate glycan chains of substrates along their passage through the Golgi apparatus (Berger and Rohrer, 2008). Mammalian glycosidases and glycosyltransferases have been demonstrated to localize in the Golgi apparatus in specific order along the cis–trans axis with some overlap, and this distribution is thought to mirror the order of glycosylation steps (Igdoura et al.,

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1999; Nilsson et al., 1993; Rabouille et al., 1995). Altering the intra-Golgi localization of glycosyltransferases has been reported to affect glycosylation pattern of the secreted reporter glycoprotein (Grabenhorst and Conradt, 1999), which indicates that the ordered distribution of the enzymes contributes to correct glycosylation. The mechanisms to achieve such subcompartmentalization of the Golgi resident glycosylation enzymes have been intensively studied in mammalian and yeast cells. Most of the glycosylation enzymes are type II integral membrane proteins comprised of a short N-terminal cytoplasmic tail, a single transmembrane domain, a stem region, and a catalytic domain oriented into the Golgi lumen (Berger and Rohrer, 2008; Schoberer and Strasser, 2011; Tu and Banfield, 2010). The information of Golgi localization is in general located within the cytoplasmic tail, the transmembrane domain, and the stem region (CTS), while the localization mechanisms can vary from enzyme to enzyme (Tu and Banfield, 2010). For some of them, the length of the transmembrane domain seems to be important (Munro, 1995). For others, not the length but the composition of amino acids of the transmembrane domain, which promotes homo- or heterooligomerization, are significant for enzyme localization (Tu and Banfield, 2010). Such heterooligomerization of the glycosylation enzymes is also suggested to ensure sequential modification of the substrates (de Graffenried and Bertozzi, 2004). Still others, especially specific yeast glycosyltransferases, are suggested to have consensus sequences in the cytoplasmic tails to associate with other proteins that also bind to COPI coats, which promotes segregation of the enzymes into transport vesicles and contributes to their localization (Moremen et al., 2012; Schmitz et al., 2008; Tu and Banfield, 2010; Tu et al., 2008). Clearly, there is no single conserved way, and moreover, these hypothetical mechanisms can act in combination. This question about the Golgi enzyme subcompartmentalization mechanism cannot be independent from the big debate about intra-Golgi trafficking (see Section 4.1), and thus is still under controversy. Rat a-2,6-sialyltransferase (ST) expressed in plant cells exhibits the localization in the trans-Golgi cisternae, very similarly to its original host organism (Wee et al., 1998), and this localization is sufficiently accomplished only by its CTS region (Boevink et al., 1998). Thus, the mechanisms of localization of the glycosylation enzymes appear to be (at least partially) conserved between animals and plants. Indeed, the fluorescence-tagged version of the CTS region of rat ST is one of the most commonly used trans-Golgi membrane marker in plant cells now (Fig. 6.2). Furthermore, the sugar chain


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structure of glycoproteins can be manipulated by swapping the CTS regions of introduced glycosyltransferases with different enzymes, which presumably alter the localization of the chimera enzymes to that of the donor of the CTS region (Bakker et al., 2006; Strasser et al., 2009). Cloning of plant glycosylation enzymes began to catch up mammals and yeast (Boisson et al., 2001; Liebminger et al., 2009; Strasser et al., 1999, 2000, 2006, 2007), which has greatly facilitated studies on their intra-Golgi localization and trafficking by live imaging. Mechanisms to locate different enzymes are proposed similarly to mammals: depending on the length of transmembrane domain (Saint-Jore-Dupas et al., 2006) or homo- or heterodimerization through CTS regions (Schoberer et al., 2013). Although ER-to-Golgi export signals are found in the cytoplasmic tails of plant Golgi membrane proteins (Hanton et al., 2005; Schoberer et al., 2009; Yuasa et al., 2005), it is not clear whether they contribute to intra-Golgi localization.

2.3.2 N-linked glycosylation Among the posttranslational modifications on proteins, N-linked glycosylation, synthesis and processing of oligosaccharides on asparagine residues in Asn-X-Ser/Thr (X represents any amino acid residues) motifs, has been most extensively studied. In all eukaryotes, N-glycosylation is initiated in the ER by en bloc transfer of the preformed oligosaccharide (Glc3Man9GlcNAc2) to the nascent polypeptide chain during translation. Next, two glucose residues are cleaved off from the N-glycan by ER a-glucosidase I and II (GCSI and GCSII) to make Glc1Man9GlcNac2, which is recognized by the ER-resident lectin-like chaperones, calreticulin (CRT), and calnexin (CNX). Subsequently, the terminal glucose residue is trimmed and the glycoprotein is freed from CRT/CNX. However, if the protein is not properly folded, its N-glycan is reglucosylated and caught by CRT/CNX, resulting in retention in the ER or retrieval back from the Golgi. Such CRT/CNX-glycoprotein binding and liberation cycle continues until the protein attains its native structure (CRT/CNX cycle) (Caramelo and Parodi, 2008). Such function of N-glycosylation in the ER quality control (ERQC) is conserved among most eukaryotes. In Arabidopsis, null mutants of glycosylation enzymes in the early steps in N-glycosylation such as GCSI and GCSII are known to be lethal (Boisson et al., 2001; Burn et al., 2002). Recently, the N-glycan dependent ERQC is suggested to be required for proper function of EFR, a pattern recognition receptor in the plant immune system for the bacterial elongation

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factor-Tu (Farid et al., 2013; Liu and Howell, 2010; Saijo, 2010). EFR is thought to be affected by N-glycosylation in protein stability, accumulation, and ligand binding (Farid et al., 2013; Ha¨weker et al., 2010). Some other plant proteins have also been reported to need N-glycans for their catalytic activity, thermostability, subcellular localization, and secretion, implying contribution of N-glycan dependent ERQC to their proper structures (Ceriotti et al., 1998; Lige et al., 2001). After they clear the ERQC system, glycoproteins go forward to the Golgi apparatus, where their high-mannose-type N-glycans acquire further modifications to become complex-type N-glycans. In mammalian cells, complex N-glycans are thought to play many important roles. For example, deletion of N-acetylglucosaminyltransferase (GnTI), the glycosyltransferase thought to initiate N-glycan processing in the Golgi, causes deficiency in complex N-glycan formation and leads to embryonic lethality in mice (Ioffe and Stanley, 1994; Metzler et al., 1994). On the other hand, mice lacking a-mannosidase II (aM-II), an enzyme acts in the early stage of Golgi N-glycan processing just after GnTI, develop an autoimmune disease (Paulson, 2007). However, in spite of the conservation of N-glycan functions in the ER between animals and plants, very little is known about the role of N-glycan processing in the plant Golgi apparatus. It is because no physiological phenotype was reported for the mutants of Golgi N-glycan modification until recently. Arabidopsis mutants defective in the Golgi glycosylation enzymes including GnTI and aM-II cannot produce normal complex N-glycans, but exhibit no growth or morphological defect under normal environment (Strasser et al., 2004, 2006; von Schaewen et al., 1993). For EFR, which requires N-glycosylation in the ER, Golgi-type modifications are dispensable (Ha¨weker et al., 2010). Artificial suppression of complex N-glycan processing by RNAi of a1,3-fucosyltransferase (FucT) and b1,2-xylosyltransferase (XylT) in tobacco and duckweed does not show obvious phenotype (Cox et al., 2006; Strasser et al., 2008). In 2008, it was reported that A. thaliana mutants defective in Golgi N-glycosylation are highly sensitive to salt stress (Kang et al., 2008). Genetic analysis indicates that the function of plasma membrane-bound endo-b1,4-glucanse KORRIGAN 1 (KOR1), which is essential for cellulose biosynthesis of cell wall, depends on complex N-glycans. Moreover, the weak mutant of KOR1 is also salt sensitive. Therefore, complex N-glycans are suggested to be necessary for sufficient cell wall formation under salt stress (Kang et al., 2008). Further investigations especially about biotic and abiotic stress responses will provide more understanding of the functions of N-glycan processing in the plant Golgi apparatus.


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2.3.3 Cell-wall polysaccharide synthesis In addition to processing of sugar chains of glycoproteins, the Golgi apparatus in plant cells plays a crucial role in synthesis of noncellulosic cell wall polysaccharides. The primary cell wall of eudicots consists of cellulose microfibrils cross-linked by hemicellulose xyloglucan (XyG), and noncellulosic polysaccharides and proteins as a matrix filling around the XyG–cellulose network. Major noncellulosic polysaccharides in the primary wall except XyG are pectic polysaccharides; homogalacturonan (HG), rhamnogalacturonan I and II (RG-I and RG-II). Unlike the polysaccharide chains on glycoproteins initially synthesized in the ER, the noncellulosic cell wall polysaccharides are assembled exclusively in the Golgi apparatus (Driouich et al., 2012). Using antibodies recognizing either the XyG backbone or the side chains, XyG has been shown to appear in trans-Golgi and the TGN (Moore et al., 1991; Zhang and Staehelin, 1992). By a similar immunocytochemical approach, HG is suggested to be synthesized in unesterified form in cis and medial cisternae followed by methylesterification in medial and trans cisternae (Moore et al., 1991; Zhang and Staehelin, 1992). For RG-I, backbone structure is synthesized similarly to HG, and arabinose-containing side chains are supposed to be added in trans cisternae (Zhang and Staehelin, 1992). Information about localization and assembly of RG-II is still missing. Most of the enzymes involved in plant cell wall polysaccharide synthesis are type II membrane proteins, albeit some are multipass transmembrane proteins (Oikawa et al., 2013). Similarly to the N-glycan glycosyltransferases, the enzymes that add side chains to XyG have been shown to localize in different Golgi cisternae in the order they suggested to work in tobacco BY-2 clls (Chevalier et al., 2010). Furthermore, the understanding about protein complex formation has recently made significant advance by application of fluorescent imaging techniques such as bimolecular fluorescence complementation (BiFC), split luciferase complementation assay (SLCA), and Fo¨rster resonance energy transfer (FRET). Using these techniques, it is becoming clear that the protein–protein interaction of glycosylation enzymes to make complexes is crucial for cell wall polysaccharide synthesis as N-glycosylation (Oikawa et al., 2013).

3. ER–GOLGI TRANSPORT 3.1. COPI vesicle 3.1.1 COPI coat (coatomer) From cell-free reconstitution assays and cell fusion experiments to monitor the cargo transport between different mammalian Golgi complexes, original

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prediction was that intra-Golgi (especially anterograde) transport was achieved by vesicle mediated trafficking (Rothman, 1994). In 1986, de novo formation of nonclathrin-coated vesicles from isolated mammalian Golgi was shown. A subset of their coat proteins were identified and shown to make a complex (coatomer), and given the name coat proteins (COPs; Serafini et al., 1991; Waters et al., 1991). This “COP-coated vesicle” was subsequently renamed COPI vesicle when the existence of COPII vesicle was established (Barlowe et al., 1994). Now, it is known that coatomer is a heptameric complex consisting of a, b, b0 , g, d, e, and z subunits (Hsu et al., 2008; Popoff et al., 2011). They are classified into two subcomplexes; one complex composed of b-, g-, d-, and z-COP forms the inner core that binds cargo, and the other complex composed of a-, b0 -, and e-COP forms the outer lattice (Popoff et al., 2011). COPI machinery is well conserved among eukaryotes including plants. All the COPI components have been found to be encoded in the Arabidopsis genome (Bassham et al., 2008). Plant COPI vesicles were first visualized by immunoelectron microscopy in Arabidopsis and maize cells by antibodies against Arabidopsis g-COP and maize d- and e-COP (Pimpl et al., 2000). Using these antisera, plant coatomers have been detected on the cisternal rims of the Golgi apparatus and the vesicles including buds formed from the Golgi (Donohoe et al., 2007; Pimpl et al., 2000; Ritzenthaler et al., 2002). In the human genome, two distinct isoforms of g-COP (g1 and g2) and z-COP (z1 and z2) have been found (Blagitko et al., 1999; Futatsumori et al., 2000). These coatomer isoforms have been demonstrated to show different intra-Golgi localization, which indicates different budding sites or functions for each of them (Moelleken et al., 2007). Arabidopsis also codes multiple isoforms of COPI proteins except for g- and s-COP; two for a-, b-, e-COP, and three for b0 - and z-COP so far (Bassham et al., 2008; Robinson et al., 2007). Whether they localize and act differently is yet to be elucidated, but it has been reported by electron tomography that there are two types of COPI vesicles (COPIa and COPIb) in Arabidopsis and scale alga Scherffelia dubia according to their coat architecture, coat thickness, and cargo staining. Moreover, COPIa vesicles bud exclusively from cis-cisternae rims and occupy the space between cis-Golgi and the ER, while COPIb vesicles bud from medial- and trans-Golgi and are distributed around these cisternae. This may indicate that COPIa vesicles are involved in the transport between the ER and the Golgi, whereas COPIb vesicles function in intraGolgi transport (Donohoe et al., 2007, 2013).


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3.1.2 ARF1 COPI vesicle formation is regulated by a GTPase ADP-ribosylation factor 1 (ARF1). Like other small GTPases, ARF1 cycles between GDP-bound inactive cytosolic state and GTP-bound active membrane-anchored state. ARF1 has hydrophobic and myristoylated N-terminus, which is exposed and inserted into the lipid bilayer by a conformational change coupled with activation by nucleotide exchange (Antonny et al., 1997; Goldberg, 1998). Activation of ARF1 by GTP binding is necessary for coatomer recruitment on the Golgi membranes (Donaldson et al., 1992a). In plant cells, ARF1 localization was observed on the Golgi and COPI vesicle membranes by immunoelectron microscopy (Pimpl et al., 2000), and later the localization to the Golgi stacks was confirmed by immunofluorescence (Ritzenthaler et al., 2002) and live imaging of fluorescence-tagged ARF1 (Stefano et al., 2006; Xu and Scheres, 2005). It is noteworthy that the immunofluorescence and ARF1-GFP signals are observed as doughnut-like rings, which indicates that ARF1 localizes especially to the Golgi cisternal rims where COPI vesicles are supposed to bud (Ritzenthaler et al., 2002; Xu and Scheres, 2005). In addition to the Golgi stacks, it has been demonstrated that ARF1 localizes to the TGN by immunofluorescence (Paciorek et al., 2005; Tanaka et al., 2009) and ARF1-GFP (Tanaka et al., 2009; Xu and Scheres, 2005), and recently by immunoelectron microscopy (Stierhof and El Kasmi, 2010). Such ARF1 population is thought to regulate the formation of clathrin-coated vesicles from the TGN (Memon, 2004; Robinson et al., 2011). Surprisingly, it has been reported that barley ARF1 expressed in onion cells colocalizes with AtARA7, the marker of late endosomes/multivesicular bodies (MVBs; Bo¨hlenius et al., 2010). Plant TGN has another face as the early endosome (see Section 6), and the TGN is recently suggested to mature into MVB along the endocytic pathway. In this context, because the compartments with an intermediate nature between the TGN and MVB are observed when endocytosis is highly activated (Choi et al., 2013), the colocalization of barley ARF1 and AtARA7 might reflect such intermediate compartments. 3.1.3 ARF GEF and GAP For activation of ARF1, guanine nucleotide exchange factors (GEFs) are needed, and they are characterized by a conserved Sec7 catalytic domain ( Jackson and Casanova, 2000). Among mammalian ARF GEFs, GBF1 is proposed to regulate COPI recruitment on the cis-Golgi and BIGs are

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specialized for clathrin-coated vesicle formation at the TGN, while the others act at post-Golgi compartments (Manolea et al., 2008; Popoff et al., 2011). ARF1-GDP meets GBF1 on the membrane to be ARF1GTP, and recruits multiple effectors including coatomers, cargo proteins, lipid-modifying enzymes, and ARF GTPase-activating proteins (GAPs; Hsu et al., 2008; Popoff et al., 2011). Brefeldin A (BFA), an antibiotic compound produced by fungi, was known to cause rapid distribution of Golgi proteins into the ER and disappearance of the Golgi apparatus in mammalian cells (Lippincott-Schwartz et al., 1989). Subsequently, BFA was found to inhibit activation step of ARF1 (Donaldson et al., 1992b; Helms and Rothman, 1992). BFA is hypothesized to bind between ARF1-GDP and the Sec7 domain of the GEF and stabilize abortive ARF1-GDP:BFA:Sec7 domain complex, which leads to the inhibition of nucleotide exchange of ARF1 and finally dissociates ARF1 and COPI coat from the Golgi membranes (Chardin and McCormick, 1999). Among ARF GEF subgroups, only GBF and BIG types are found in plants (Teh and Moore, 2007). The first ARF GEF identified in plants was the Arabidopsis GBF protein GNOM (GN). Although animal and fungal GBF proteins act at the Golgi, GNOM was revealed to function mainly in the recycling process of auxin efflux carrier PIN1 from endosomes to the plasma membrane (Geldner et al., 2003). Later, GNOM-LIKE1 (GNL1), the closest homolog of GN in Arabidopsis, was identified and shown to be responsible for ER–Golgi trafficking. GNL1 primarily localizes to the Golgi stacks and regulates coatomer recruitment there (Richter et al., 2007). Because GN can functionally substitute for GNL1 but not vice versa, GN is supposed to gain the plant-specific function in addition to the ancestral role of GBF proteins in ER–Golgi trafficking (Richter et al., 2007). Since GNL1 is BFA resistant, BFA treatment does not cause Golgi disruption in Arabidopsis root cells unlike in mammalian and tobacco cells (Teh and Moore, 2007). In order to fuse with target membranes, coated vesicles need to be uncoated. For COPI vesicle uncoating, inactivation of ARF1 is necessary, which requires GAPs (Bigay et al., 2003; Melançon et al., 1987; Reinhard et al., 2003). However, the ARF GAP activity was revealed necessary not only for vesicle uncoating but also for COPI cargo sorting and vesicle formation (Lee et al., 2005; Nie and Randazzo, 2006). The GAP activity of mammalian ARF GAP1 is accelerated by binding with coatomer, but this effect is inhibited by a p24 protein, which is suggested to act as a cargo


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receptor (Goldberg, 2000). Therefore, ARF GAP1 is presumed to act in “proofreading” of COPI vesicle formation, because only the coatomers with cargo proeins are thought to stay on the membrane long enough to be polymerized. Moreover, it has been also reported that ARF GAP1 activity increases as the membrane curvature approaches that of a typical transport vesicle, which might ensure vesicle uncoating after completion of vesicle formation (Bigay et al., 2003). In plant cells, although the Arabidopsis genome encodes 15 ARF GAP domain (AGD) proteins, most of their functions are unknown. AGD7, the closest homolog of human ARF GAP1, has been revealed to localize to the Golgi apparatus and regulate ARF1 activity in ER–Golgi trafficking (Min et al., 2007). In addition, AGD8 and AGD9, homologs of Glo3p-type ARF GAPs functioning in COPI vesicle formation in animal and yeast cells, are reported to recruit ARF1-GDP from the cytosol to the Golgi membranes (Min et al., 2013). Such a role has not been reported for mammalian or yeast ARF GAPs. 3.1.4 Golgi-to-ER transport Although COPI vesicles were thought to act in the anterograde trafficking in the early studies, this initial view was changed by subsequent findings indicating their roles in Golgi-to-ER retrograde transport. Soluble ER proteins have a common KDEL or HDEL motif in their C-termini in animal and yeast cells, and this motif is recognized by the receptor coded by yeast ERD2 gene and its homologs in animals. Erd2 captures leaked ER luminal proteins mainly at the cis-Golgi or ER-Golgi intermediate compartment (ERGIC), and returns them to the ER (Lewis and Pelham, 1990; Lewis et al., 1990; Semenza et al., 1990). This Golgi-to-ER transport step has been shown to depend on COPI vesicles (Girod et al., 1999; Lewis and Pelham, 1996). For transmembrane proteins, ER retrieval signals in the cytosolic domain are known to bind directly to coatomer subunits (Letourneur et al., 1994; Michelsen et al., 2007). In yeast cells, a subset of ER membrane proteins are recognized through their transmembrane region by Rer1 at the Golgi and retrieved to the ER, and this process also depends on COPI vesicles (Sato et al., 1995, 2001, 2003). These recognition systems for Golgi-to-ER retrograde traffic by COPI vesicles are also conserved in plants. C-terminal KDEL and HDEL sequences are sufficient for ER localization of soluble proteins (Denecke et al., 1992). The Arabidopsis homolog of ERD2 can complement the yeast erd2 deletion mutant (Lee et al., 1993), and localizes to the Golgi stacks in

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Arabidopsis and tobacco cells (Boevink et al., 1998; Saint-Jore et al., 2002; Takeuchi et al., 2000). There are three RER1 homologs in Arabidopsis (RER1A, RER1B, and RER1C) and all can complement yeast rer1 deletion in different degrees with RER1B being the best (Sato et al., 1999). GFPRER1B has been demonstrated to localize to the cis-Golgi in Arabidopsis and tobacco cultured cells (Ito et al., 2012; Takeuchi et al., 2000). ER-localized proteins with the COPI-interacting ER retrieval signals have been also identified (Benghezal et al., 2000; Boulaflous et al., 2009; Contreras et al., 2004; McCartney et al., 2004; Montesinos et al., 2012). Because BFA inhibits the formation of COPI vesicles responsible for Golgi-to-ER transport, it seems puzzling that the secretion is disrupted and the Golgi enzymes relocalize to the ER by BFA treatment in many cell types. In yeast, mutation in UFE1, which encodes the SNARE protein necessary for COPI vesicle fusion with the ER, exhibits blockade of secretion and ER retention of a vacuolar protein (Lewis and Pelham, 1996). In plant cells, overexpression of GDP-locked dominant negative form of ARF1 (ARF1T31N) causes absorbance of ERD2 and CTS regions of ST and XylT into the ER, and inhibits secretion of a-amylase (Stefano et al., 2006; Takeuchi et al., 2002). Moreover, GTP-locked constitutive active ARF1 mutant protein (ARF1Q71L) causes accumulation of ERD2 and a vacuolar protein sporamin in the ER (Takeuchi et al., 2002). Because weak expression of this construct only inhibits Golgi-to-ER transport (Langhans et al., 2008), blockade of ER export is supposed to be a secondary effect. From these facts, the depression of ER-to-Golgi anterograde trafficking by BFA treatment is thought to be the indirect consequence of the inhibition of the retrograde trafficking, which is presumably necessary for salvaging components for COPII vesicle formation (such as SNAREs). However, the disappearance of the Golgi apparatus cannot be explained only by disruption of transport between the ER and the Golgi. Actually in mammalian cells, it has been reported that the Golgi apparatus does not disappear by combining a cationic ionophore monensin with BFA, in spite of the dissociation of ARF1 and coatomer from the Golgi membranes and the blockade of ER export (Barzilay et al., 2005). In tobacco BY-2 cells, hybrid structures of the ER and the Golgi stacks have been observed upon BFA treatment by electron microscopy, which suggested direct fusion of the two organelles (Ritzenthaler et al., 2002). The absorption of the Golgi proteins to the ER was proposed to result from this fusion. It was hypothesized that this phenomenon was due to Golgi accumulation of the SNARE proteins responsible for fusion of Golgi-derived COPI vesicles to the ER, although there is no experimental support. Anyway, these data indicate that the


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existence of the Golgi apparatus depends on the delicate balance between anterograde and retrograde trafficking between the ER and the Golgi. COPI vesicles are also believed to be responsible for intra-Golgi trafficking. About this transport pathway, we will describe in Section 4. 3.1.5 Tethering factors and SNAREs The fundamental process of membrane fusion is mediated by long-range tethering factors mostly regulated by RAB GTPases, and short-range SNARE proteins (Saito and Ueda, 2009; Szul and Sztul, 2011). For the Golgi-to-ER COPI vesicles, tethering to the ER membrane is mediated by Dsl1 trimeric complex in yeast cells (Ren et al., 2009). Dsl1 complex is composed of Dsl1, Tip20, and Dsl3/Sec39, and Dsl1 binds to a- and d-COP (Andag and Schmitt, 2003; Reilly et al., 2001). Dsl1 complex stably localizes to the ER membrane independently to any RAB GTPases (Sztul and Lupashin, 2009). Membrane fusion is mediated by ER-localized Q-SNAREs (Qa: Ufe1, Qb: Sec20, and Qc: Use1) and R-SNARE Sec22 (Ballensiefen et al., 1998; Burri et al., 2003; Dilcher et al., 2003; Lewis and Pelham, 1996). Dsl1 complex interacts with the SNARE complex and is suggested to promote SNARE complex assembly during membrane fusion (Kraynack et al., 2005; Ren et al., 2009). In mammalian cells, the homologous complex of Dsl1 composed of ZW10, RINT-1, and NAG has been identified and also interacts with ER Q-SNAREs (Schmitt, 2010; Sztul and Lupashin, 2009). Among three subunits of Dsl1 complex, a homolog of Tip20 was first identified in Arabidopsis. It was isolated as the gene responsible for maigo2 (mag2) mutant, which accumulates seed storage proteins in the ER. MAG2/ AtTIP20 protein is associated with the ER membrane, and interacts with SNAREs AtSEC20 and AtSYP81/AtUFE1 (Li et al., 2006). CFP-tagged MAG2/AtTIP20 expressed in tobacco leaf epidermal cells has been shown to colocalize with the Golgi stacks and move together (Lerich et al., 2012). A recent study identified three MAG2/AtTIP20 interators designated MAG2 INTERACTING PROTEINS (MIPs,) and the mutant plants lacking any one of them developed abnormal structures with storage proteins in the ER of seed cells. Because MIP1 and MIP2 were revealed to have a ZW10 domain and a Sec39 domain, respectively, the MAG2/AtTIP20– MIP1–MIP2 complex is suggested to be homologous to Dsl1 complex. However, this complex has the fourth subunit, MIP3, which contains a Sec1 domain. This is the first report of a Sec1-containing protein involved in ER–Golgi trafficking including animals and yeast (Li et al., 2013).

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By live-cell imaging using GFP-tagged proteins, six SNAREs are shown to localize to the ER in plant cells: Qa-SNARE AtSYP81/AtUFE1; Qc-SNARE AtSYP71, AtSYP72, and AtSYP73; and R-SNARE AtSEC22 and AtVAMP723 (Uemura et al., 2004). No Qb-SNARE was found on the ER membrane in this study, but AtSEC20 is suggested to localize to the ER. In addition, AtUSE1 was recently reported to localize to the ER (Lerich et al., 2012). Since overexpression of AtSYP81/AtUFE1 causes relocalization of Golgi enzymes, it is thought to function in the transport between the ER and the Golgi (Bubeck et al., 2008). SYP7 family is not conserved in yeast and animals, and its involvement in COPI vesicle transport is not clear (Sanderfoot et al., 2000). However, AtSYP72 tagged by fluorescent protein marks punctate signals along the ER tubules, and AtSYP81/AtUFE1 and COPI dependent Golgi-to-ER cargo proteins colocalize at the puncta only when coexpressed with AtSYP72. Such punctate signals do not move with the Golgi stacks, but when the Golgi stacks are immobilized by actin depolymerization, they colocalize with the Golgi signals. Therefore, it is proposed that Golgi-to-ER retrograde vesicles are tethered by Dsl1 complex while the Golgi stack is moving, and fuse to the ER during temporary pauses of the Golgi on the AtSYP72 puncta (Lerich et al., 2012).

3.2. COPII vesicle and ERES 3.2.1 COPII vesicle formation COPII coat assembly in S. cerevisiae is initiated by activating small GTPase Sar1 by its GEF, Sec12 (Barlowe and Schekman, 1993; Nakano and Muramatsu, 1989). Sec12 has been revealed to interact with Rer1 through its transmembrane domain, therefore, Sec12 is regulated to localize to the ER membrane by Golgi-to-ER COPI vesicles, which results in restriction of Sar1 activation on the ER (Sato et al., 1996). Sar1 has a hydrophobic a-helix at the N-terminus similarly to one of its closest evolutionary relatives Arf1, and is believed to associate with the membrane by embedding the N-terminal domain into the bilayer upon activation (Bi et al., 2002; Huang et al., 2001). In contrast to heptameric coatomer, which is recruited to the membrane en bloc, COPII coat complexes (Sec23/24 and Sec13/31) are recruited sequentially to form layers. Activated Sar1 recruits Sec23/24 complex first by interacting with Sec23 subunit to form Sec23/24-Sar1 “prebidding complex” (Bi et al., 2002; Yoshihisa et al., 1993). The prebudding complexes recognize cargo proteins and compose the inner layer of COPII coat (Kuehn et al., 1998). The assembly of the prebudding


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complex is stabilized through interactions with transmembrane cargo proteins, and different kinetics between the disassembly of the prebudding complex and the cycle of Sar1-GTP hydrolysis facilitates proper and efficient cargo sorting and vesicle formation (Sato and Nakano, 2005). The outer layer is composed of Sec13/31 complex organized into a cage (Matsuoka et al., 2001). Sec13/31 complex interacts with the prebudding complex through binding of Sec31 to Sec23 and Sar1 (Bi et al., 2007). The outer cage assembly is likely to collect the prebudding complexes and drive membrane deformation. COPII components are highly conserved in eukaryotes. Notably, the Arabidopsis genome encodes multiple isoforms for each COPII component (two Sec12, five Sar1, seven Sec23, three Sec24, two Sec31, and two Sec31; Robinson et al., 2007). Among them, mutant plants have been reported only for Sec24 isoforms. A recessive missense point mutation in AtSEC24A, which causes partial retention of Golgi and secretory proteins in the abnormally shaped ER, has been independently identified by two groups by forward genetics combined with visual screening using Golgi or ER markers. Moreover, Atsec24A null mutant was revealed to be lethal, indicating that AtSEC24A is essential (Faso et al., 2009; Nakano et al., 2009). Recently, mutants of AtSEC24B and AtSec24C were reported, although the organelles and intracellular transport is yet to be investigated in detail (see Section 3.2.2; Tanaka et al., 2013). The functions of the plant homologs of Sec12 and Sar1 have been confirmed in alternative ways. AtSEC12 suppresses the yeast sec12 null mutant (d’Enfert et al., 1992). In addition, overexpression of AtSEC12 in tobacco protoplasts inhibits secretion, and this effect is rescued by additional overexpression of AtSAR1B (Phillipson et al., 2001). AtSAR1B and tobacco (Nicotiana tabacum) SAR1 also have been revealed to suppress yeast sar1 null mutant and temperature-sensitive sec12 or sec16 mutants, which indicates the functional conservation (d’Enfert et al., 1992; Takeuchi et al., 1998). The GTP-locked dominant mutant protein of AtSAR1B causes retention of Golgi and vacuolar proteins in the ER and inhibits secretion (Phillipson et al., 2001; Takeuchi et al., 2000). GTP-locked AtSAR1A is also reported to inhibit secretion but to a less extent than AtSAR1B, and the membrane binding affinity of AtSAR1B is stronger than AtSAR1A (Hanton et al., 2008). Whether such differences reflect functional difference is not known yet. Similarly, at least two isoforms of N. tabacum SAR1 proteins inhibits ER export in their GTP- or GDP-locked forms (Andreeva et al., 2000; daSilva et al., 2004; Osterrieder et al., 2010; Robinson et al.,

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2007; Schoberer et al., 2009; Yang et al., 2005). The expression of GDPlocked form of one of the N. tabacum SAR1s in BY-2 cells causes dissociation of SEC13 from the ER membrane (Yang et al., 2005). Recently, gene knockdown by RNAi of three rice SAR1 homologs specifically in the endosperm was shown to block ER export of seed storage proteins (Tian et al., 2013). The presence of COPII vesicles in plant cells is not fully established. Although there are a few reports finding vesicle-like structures by electron microscopy (Donohoe et al., 2007, 2013; Langhans et al., 2012; Staehelin and Kang, 2008), such structures are rarely found especially in highly vacuolated cells (Brandizzi and Barlowe, 2013; Hawes, 2012). This is thought to be due to rapid consumption of the vesicles at the ER–Golgi interface. It is impossible to rule out the possibility that ER–Golgi transport is mediated by direct connection by tubules; however, there are too few reports supporting this idea (Brandizzi et al., 2002; Hawes, 2012).

3.2.2 Cargo sorting During COPII vesicle formation, the prebudding complex interacts with cargo proteins and concentrates them in the budding vesicle (Kuehn et al., 1998). The role of Sec24 in cargo recognition has been revealed by structural, biochemical, and genetic approaches using yeast (Miller et al., 2003; Mossessova et al., 2003). As described above, there are three SEC24 isoforms in the Arabidopsis genome. Because Atsec24A null mutant is lethal, and the aberrant morphology of the ER and the Golgi caused by the point mutation in AtSEC24A cannot be suppressed by overexpression of AtSEC24B or AtSEC24C, these isoforms might be functionally different (Faso et al., 2009; Nakano et al., 2009). A further study showed that the loss of AtSEC24A function induces a defect in pollen germination but does not affect the female gametophyte (Conger et al., 2011). On the other hand, AtSEC24B and AtSEC24C were shown to function redundantly in both male and female gametogenesis (Tanaka et al., 2013). Moreover, microarray analysis of microspores indicates that the AtSEC24A expression peak comes earlier than that of AtSEC24B during pollen development (Conger et al., 2011). Since Arabidopsis SEC24 proteins are also thought to function in cargo sorting like the homologs in yeast, the difference between AtSEC24A and AtSEC24B/C may reflect specific cargo recognition. Indeed, such differentiation among Sec24 isoforms in cargo recognition has been reported for


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mammals and yeast (Kurihara et al., 2000; Merte et al., 2010; Pagano et al., 1999; Roberg et al., 1999; Wendeler et al., 2007). The particular recognition targets of AtSEC24 isoforms are yet to be characterized. 3.2.3 Uncoating, tethering, and fusion Unlike the COPI coat, Sec23, a subunit of the COPII coat complex, is known to act also as the GAP for Sar1 (Bi et al., 2002; Yoshihisa et al., 1993). In addition to the basic GAP activity of Sec23, Sec31 binding to both Sec23 and Sar1 accelerates Sar1-GTP hydrolysis by an order of magnitude (Antonny et al., 2001; Bi et al., 2007). Sar1 inactivation rapidly leads to COPII coat disassembly (Antonny et al., 2001). GTP hydrolysis is not required for COPII budding itself (Barlowe et al., 1994; Oka and Nakano, 1994) and was thought to be dispensable for cargo loading (Barlowe et al., 1994), but now it is known to contribute to efficient and proper cargo selection as described in Section 3.2.1. There are seven SEC23 isoforms in the Arabidopsis genome, greatly outnumber the only one Sec23 in yeast and two in human (Robinson et al., 2007). The localization of one of the AtSEC23s has been observed by immunofluorescence (Yang et al., 2005) and another by fluorescent protein tagging (see Section 3.2.4; Stefano et al., 2006), and the latter has been shown to bind AtSEC24A (Stefano et al., 2006). Both are believed to function similarly to the yeast homolog. The functional significance of AtSEC23 multiplicity is unknown. As the tethering factors for COPII vesicles, a long coiled-coil protein p115 (yeast Uso1) and a multisubunit complex transport protein particles I (TRAPPI) are well known. p115 and yeast homolog Uso1 have been revealed to be essential for COPII tethering. In the case of Uso1, it is thought to tether COPII vesicles directly to the cis-Golgi (Cao et al., 1998; Sapperstein et al., 1996). On the other hand, the target of COPII vesicle tethering by p115 is less clear in mammalian cells; in addition to the cis-Golgi cisternae, it can be other COPII vesicles and ERGIC (Allan et al., 2000; Szul and Sztul, 2011; Xu and Hay, 2004). p115 and Uso1 are recruited to COPII vesicles by the active form of the small GTPases Rab1 and its yeast homolog Ypt1, respectively (Allan et al., 2000; Cao et al., 1998). From yeast studies, TRAPPI complex is known to act as the GEF for Ypt1 (Wang et al., 2000). TRAPPI itself also interacts directly with Sec23 subunit through Bet3 subunit and function as a multisubunit tether (Cai et al., 2007; Sacher et al., 1998). In plants, homologs of p115 and all TRAPPI subunits have been found (Latijnhouwers et al., 2005). Arabidopsis p115 homolog was first isolated as

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GOLGIN CANDIDATE 6 (GC6), and the GFP-fusion protein or only the C-terminal domain tagged with GFP was shown to localize to the restricted domain of the Golgi stacks (Latijnhouwers et al., 2007). The localization of GC6/Atp115 has been also observed by immunoelectron microscopy, which locates GC6/Atp115 to the surface of COPII vesicles and the ribosome-exclusion area (Golgi matrix region) of the cis-Golgi cisternae in Arabidopsis root cells and tobacco BY-2 cells (Kang and Staehelin, 2008). Subsequently, the responsible gene for Arabidopsis maigo4 mutant, which accumulates seed storage proteins in the ER, was revealed to be GC6/Atp115 (Takahashi et al., 2010). The function of the Arabidopsis homolog of TRAPPI complex is yet to be investigated. The function of Ypt1/Rab1 is likely to be conserved also in plants, because Rab1 homologs from some plants have been reported to complement the yeast ypt1 mutant (Kim et al., 1996; Park et al., 1994). Rab1 homologs in higher plants are called RABD group, and divided into two subclasses, RABD1 and RABD2, according to their sequences (Pinheiro et al., 2009). Arabidopsis has one RABD1 and three RABD2 (RABD2a, RABD2b, and RABD2c), and the RABD2 members are thought to be extensively redundant. Both subclasses are involved in ER–Golgi trafficking because dominant negative forms of RABD1 or RABD2a inhibit ER export of secretory cargo and some Golgi or vacuolar proteins (Batoko et al., 2000; Park et al., 2004; Pinheiro et al., 2009; Saint-Jore et al., 2002; Samalova et al., 2006; Zheng et al., 2005). However, the dominant negative effects of a mutant protein of one subclass cannot be suppressed by overproduction of the wild-type protein of the other subclass, suggesting that these subclasses are functionally different. In support of this, RABD2 subclass is essential but RABD1 subclass is not. Nevertheless, rabD2b rabD2c double mutants are normal but rabD1 rabD2b rabD2c triple mutants are short and bushy with low fertility, indicating that these two subclasses have partially overlapping functions (Pinheiro et al., 2009). For membrane fusion of COPII vesicles to the cis-Golgi after tethering, the SNARE proteins Sed5 (Qa), Bos1 (Qb), Bet1 (Qc), and Sec22 (R-SNARE) are thought to be responsible in yeast (Barlowe and Miller, 2013). Nine Arabidopsis SNARE proteins have been localized to the Golgi apparatus by a systematic analysis using Arabidopsis cultured cells: Qa-SNARE AtSYP31 and AtSYP32; Qb-SNARE AtGOS11, AtGOS12, AtMEMB11, and AtMEMB12; Qc-SNARE AtBS14a and AtBS14b; and R-SNARE AtVAMP714 (Uemura et al., 2004). In addition, AtSEC22 expressed in tobacco leaf epidermal cells or protoplasts is shown to localize


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to the ER and the Golgi stacks (Bubeck et al., 2008; Chatre et al., 2005). AtSYP31 and AtSYP32 are Sed5 homologs, and AtMEMB11 and AtMEMB12 are homologs of membrin, the mammalian functional homolog of Bos1 (Sanderfoot et al., 2000). Overexpression of AtSYP31, MEMB11, or AtSEC22 has been reported to cause relocalization of Golgi proteins into the ER, which indicates their involvement in ER–Golgi transport (Bubeck et al., 2008; Chatre et al., 2005). AtBS14a and AtBS14b are similar to Bet1 and Sft1, another Golgi-localized Qc-SNARE from the same family of Bet1. The overexpression of either AtBS14a or AtBS14b is reported to support the growth of the yeast sft1 null mutant but not the bet1 mutant (Tai and Banfield, 2001). Therefore, AtBS14a and AtBS14b may be responsible for intra-Golgi trafficking like yeast Sft1 (see Section 4.2). 3.2.4 ERES COPII vesicles are known to bud from specialized domains of the ER membrane called transitional ER (tER) or ER exit/export sites (ERES). The existence of such domains with intermediate features between the rough ER and the Golgi vesicles was suggested long before the identification of the coated vesicles (Palade, 1975). It was later revealed that the COPII components are enriched in the ERES by immunoelectron microscopy in mammal and yeast cells, therefore, the COPII components became convenient markers for the ERES (Kuge et al., 1994; Orci et al., 1991; Paccaud et al., 1996; Rossanese et al., 1999). By immunofluorescence imaging or live imaging using fluorescent protein-tagged COPII components, the ERES appear as multiple discrete puncta (Hammond and Glick, 2000; Horton and Ehlers, 2003; Okamoto et al., 2012; Rossanese et al., 1999). Among the Sec proteins required for vesicle formation from the ER, the function of Sec16 has been long unknown. Sec16 localizes to the ERES and its inactivation causes disruption of the ERES in several species (Bhattacharyya and Glick, 2007; Connerly et al., 2005; Ivan et al., 2008; Watson et al., 2006; Yorimitsu and Sato, 2012). Sec16 is a large multidomain peripheral protein of the ER membrane, and binds to the COPII components through independent domains for each subunit (Espenshade et al., 1995; Gimeno et al., 1996; Shaywitz et al., 1997). From these facts, it has been speculated that Sec16 acts as the scaffold for COPII coat assembly at the ERES. Recently, Yorimitsu and Sato (2012) have reported that yeast Sec16 interferes with Sec31-mediated Sec23 GAP activation, which leads to stabilization of COPII coat complex and prevents premature complex

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disassembly. In addition, Sec16 alone has the ability to self-assemble into homooligomeric complexes (Yorimitsu and Sato, 2012). This Sec16 cluster is suggested to contribute as the basis of the ERES, and spatially restricts the vesicle budding. In the Arabidopsis genome, two Sec16 homologs are predicted (Robinson et al., 2007). Mammals also have two Sec16 homologs, one being about twice as long as the other. The two proteins are both necessary for normal ERES organization and ER export (Bhattacharyya and Glick, 2007). Two Arabidopsis SEC16 proteins are predicted to have almost the same size as the shorter counterpart of mammals (Robinson et al., 2007). A most recent study has revealed that one of them, SEC16A/MAG5, interacts with COPII coats and localizes to the ERES. Its mutant plants exhibit compromised ER export, indicating its role in ER-to-Golgi trafficking (Takagi et al., 2013). In the cells with discrete Golgi stacks such as P. pastoris, Caenorhabiditis elegans, and Drosophila melanogaster, the ERES and the Golgi stacks are almost always closely associated to form “secretory units” (Kondylis and Rabouille, 2009; Rossanese et al., 1999; Witte et al., 2011). In mammalian cells with the concentrated Golgi, the ERGIC is frequently associated close to the ERES instead of the Golgi complex (Appenzeller-Herzog and Hauri, 2006; Bannykh et al., 1996). Moreover, when the microtubules are disrupted by nocodazole treatment in mammalian cells, the Golgi ministacks are located near the ERES like in invertebrate cells (Hammond and Glick, 2000). Also in plant cells, many punctate signals can be observed by using COPII coat subunits as markers. However, conflicting results have been reported about the relationship between the ERES and the Golgi stacks. By immunofluorescence observation of COPII coat proteins, the signals labeled many punctate structures greatly outnumbering the Golgi stacks. Some of them were associated with the Golgi stacks, and such signals were often found on the areas of the ER tubules associated with the Golgi stacks. Furthermore, by live imaging of tomato SEC13 tagged with GFP in tobacco BY-2 cells, the signal pattern was similar to the immunofluorescence, and the Golgi stacks were observed to associate with the SEC13-GFP signal around their rims. The level of SEC13-GFP around the Golgi stacks varied along the time, and the Golgi stacks moving slower tended to have more SEC13 signals (Yang et al., 2005). GFP-tagged tobacco SAR1, which can inhibit ER–Golgi transport in the dominant negative mutant form, also labeled puncta either associated or nonassociated with the Golgi stacks


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(Robinson et al., 2007). From these data, the “kiss-and-run” model was suggested: the ERES are relatively stable, and the Golgi stacks temporarily associates with them and ER-to-Golgi transport takes place during their contacting periods (Fig. 6.4A). On the other hand, tobacco SAR1 and Arabidopsis SAR1, SEC23, SEC24, and SEC13 fused with fluorescent proteins marked puncta associated with the Golgi stacks, without signals independent from the Golgi. The COPII signals moved always with the associated Golgi stacks (daSilva et al., 2004; Hanton et al., 2007, 2008, 2009; Stefano et al., 2006). The punctate signals of YFP-SEC24 increased in number and became more definite by expression of the cargo with COPII-dependent ER export motif, indicating the involvement of the puncta of YFPSEC24 in ER export (Hanton et al., 2007). Moreover, FRAP analysis showed that ER-to-Golgi protein transport can occur while the Golgi stacks are moving (daSilva et al., 2004). Therefore, the “secretory unit” model was suggested: the Golgi stacks and the ERES are continuously associated and the ER-to-Golgi transport continues during rapid movement (Fig. 6.4B). Our recent study has demonstrated that Arabidopsis SEC13 fused with YFP expressed in tobacco BY-2 cells labeled many puncta outnumbering the Golgi stacks, and some of them are associated with the Golgi in ring-like shapes surrounding the Golgi rims. Moreover, the SEC13-YFP signal moved together with the associated Golgi stacks (Ito et al., 2012). These results suggest that the truth is in between the two extreme models (hybrid model, Fig. 6.4C). Because the SEC13-YFP puncta without associated Golgi stacks were smaller and extremely mobile independently of actin filaments, we speculate that they become stable and active when they encounter the Golgi stacks. Recent work suggested that the COPII signal is not on the ER but in the small gap between the ER and the Golgi, which represents the COPII vesicles accumulating at the cis-face of the Golgi, not the ERES (Langhans et al., 2012). In Arabidopsis leaf epidermal cells, pulling of individual Golgi stacks by a focused infra-red laser beam (laser tweezers) was shown to drag the ER membrane as tubules behind them (Sparkes et al., 2009b). This result indicates the physical connection between the Golgi and the ER, which involves Golgi matrix and tethering proteins. Such matrix proteins are thought to catch the COPII vesicles before fusion with the cis-Golgi while the Golgi stacks are moving, therefore Langhans et al. (2012) suggested a difficulty to know whether the ERES move together with the Golgi stacks or not by the observation of only COPII coats. Furthermore, COPI vesicles responsible for Golgi-to-ER transport are proposed to fuse with the ER

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Figure 6.4 Models for ERES–Golgi organization in plants cells. (A) The kiss-and-run model. In this model, the ERES are relatively stable, and the Golgi stacks travel from one ERES to another. ER-to-Golgi transport takes place during the temporal association of the Golgi stacks and the ERES. (B) The secretory unit model. In this model, the Golgi stacks and the ERES are continuously associated and move together. ER-to-Golgi transport occurs not only during their pauses but also during rapid movement. (C) The hybrid model. In this model, some of the ERES are continuously associated with the Golgi stacks similarly to the secretory unit model, but the others are not. The ERES without associated Golgi stacks are smaller and move independently from the Golgi stacks. They get stable and active when they encounter the Golgi stacks. (D) The modified secretory unit model. In this model, the temporal pauses of the Golgi stacks take place at the ERIS. The COPII and COPI vesicles are continuously formed during the Golgi movement, but their fusion with the target membranes only occurs while the Golgi stacks are immobile. Both types of vesicles accumulate between the Golgi and the ER, and move together with the Golgi stacks.


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during temporary pauses at “ER import sites” (ERIS, see Section 3.1.5; Lerich et al., 2012). These data are summarized and called as the modified secretory unit model (Fig. 6.4D; Lerich et al., 2012). Unlike the COPII coat subunits, the distribution of Sec12 on the ER membrane is different between species. Sec12 is concentrated to the ERES near the Golgi stacks in P. pastoris, whereas Sec12 is dispersed throughout the ER in S. cerevisiae (Rossanese et al., 1999). Because AtSEC12-YFP expressed in tobacco cells localizes throughout the ER, it is not appropriate as an ERES marker in plant cells (daSilva et al., 2004). Analysis using SEC16 should shed light on the controversy. The plant Golgi stacks are observed associated with the ER tubules and the rims of the ER cisternae, but not with the ER cisternal surface (Fig. 6.5; Sparkes et al., 2009a). If active ERES continuously associate and move with the Golgi stacks, the ERES should locate to the ER domains with high membrane curvature. Such curvature of the ER is at least partially regulated by reticulon family proteins (Sparkes et al., 2010). Recently, Okamoto et al. (2012) have revealed that the high curvature of the ER membrane, especially saddle-like structures that contain both positive and negative curvatures, is important for ERES localization in yeast. In the S. cerevisiae cells lacking reticulon family proteins, peripheral ER network was disrupted, and the ERES were localized to the remaining few high-curvature domains. Moreover, the dynamics and morphology of the Golgi were affected in the mutants. Similar regulation may well exist in plant cells.

Figure 6.5 Association of the ER tubules and the Golgi stacks. Confocal image of a tobacco BY-2 cell expressing SP-GFP-HDEL (ER lumen, green) and mRFP-AtSYP31 (cis-Golgi, magenta). The ER tubules seem to surround the Golgi stacks on the cisternal rims. Scale bar, 10 mm.

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3.3. ERGIC Compared to plants, fungi, and invertebrates, the Golgi apparatus is far from the ERES in mammalian cells because the Golgi is concentrated near the nucleus. ERGIC consists of vesiculotubular clusters, and is marked by a lectin ERGIC-53 (Appenzeller-Herzog and Hauri, 2006; Schweizer et al., 1990). Observations of the glycoprotein cargo transport revealed that the structure is the intermediate compartment of transport between the ER and the Golgi (Saraste and Svensson, 1991; Schweizer et al., 1990). In the ERGIC, sorting and concentration of anterograde and retrograde cargo by COPI vesicles are thought to take place (Martıńez-Mena´rguez et al., 1999). About the transport between the ER and the Golgi via the ERGIC, there are two models. In the “transport complex model,” ER-derived COPII vesicles fuse with each other to form the ERGIC, and the ERGIC travels to the Golgi on the microtubule to become the cis-Golgi by fusing with other ERGIC or to fuse with preexisting cis-Golgi. This model is supported by the observations that ERGIC moves from the ERES to the Golgi along microtubules (Presley et al., 1997; Scales et al., 1997). On the other hand, in the “stable compartment model,” ERGIC is a static structure and facilitates a two-step transport process. First, COPII vesicles mediate short-range transport from the ER to the ERGIC independently of microtubules, and next the anterograde cargo-rich domains of the ERGIC are transported long distance to the cis-Golgi depending on microtubule tracks. A live imaging study revealed that the ERES was stationary and small soluble cargo but not ERGIC-53 was exported to the Golgi (Ben-Tekaya et al., 2005). It is also possible that different mechanisms apply for different cargo proteins. In the cells whose Golgi stacks are closely associated to the ERES, the presence of ERGIC has not been presumed. However, a recent report suggests that C. elegans cells have ERGIC-like compartments between the cisGolgi and the ERES (Witte et al., 2011). Also in plant cells, it is proposed that the cis-most Golgi cisternae are biosynthetically inactive and function as the site of membrane assembly and cargo sorting, similarly to the mammalian ERGIC (Donohoe et al., 2013). Moreover, we have demonstrated by BFA treatment and its removal of tobacco BY-2 cells that punctate structures with some cis-Golgi components near the ERES act as the scaffold for Golgi stack regeneration, suggesting their ERGIC-like properties (see Section 5.4; Ito et al., 2012).


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4. INTRA-GOLGI TRAFFICKING 4.1. Models of intra-Golgi trafficking To discuss the formation of the Golgi structure, we cannot skip the long-lasting debate about intra-Golgi trafficking (Glick and Luini, 2011; Glick and Nakano, 2009; Nakano and Luini, 2010). This debate is about how cargo molecules are transported through the stack of cisternae, and has involved many cell biologists. Among many models proposed, two major models were the issue of dispute; one is the vesicular transport (stable compartments) model, and the other is the cisternal maturation model (Fig. 6.6).

Figure 6.6 Two major models for intra-Golgi trafficking. (Left) The vesicular transport (stable compartments) model. In this model, the Golgi cisternae are stable and distinct suborganelles with different resident proteins. Cargo proteins are transported from one cisterna to the next by anterograde COPI vesicles, whereas the Golgi proteins are excluded from them and remain in the cisternae. Cargo proteins reach the TGN and exit by clathrin-coated vesicles or other carriers. (Right) The cisternal maturation model. In this model, the Golgi cisternae themselves function as the cargo carriers. They are transient compartments that are newly formed by homotypic fusion of COPII vesicles, progress from cis to trans, and break down at the TGN stage. The Golgi proteins recycle back from later to earlier cisternae by retrograde COPI vesicles, and the nature of the cisternae gradually changes as they progress.

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In the vesicular transport model, the Golgi stack is viewed as a series of distinct suborganelles, each of which has a characteristic set of resident proteins. Cargo proteins travel from one cisterna to the next by anterograde vesicles, and the Golgi resident proteins remain in the cisterna by exclusion from the vesicles. The fundamental idea of this model can be traced back to the data which demonstrated that the peri-Golgi vesicles contained newly synthesized secretory proteins ( Jamieson and Palade, 1966). As described in Section 3.1.1, cell-free reconstitution assays and cell fusion experiments in 1980s indicated vesicular transport between cisternae, and subsequently identified COPI vesicles were thought to be responsible for this transport. The model was updated later in which COPI vesicles move bidirectionally, with anterograde vesicles carrying secretory cargo and retrograde vesicles recycle trafficking components (Orci et al., 2000b). On the other hand, the cisternal progression model, which was upgraded later to the cisternal maturation model, was first proposed from morphological studies (Grasse´, 1957). In this model, Golgi cisternae themselves are thought as the cargo carriers. Homotypic fusion of ER-derived COPII vesicles or other carriers like ERGIC nucleates the new cis-cisternae, which progress forward along the secretory pathway to become the TGN. The Golgi cisternae are transient compartments and continuously turn over. In the updated cisternal maturation model, COPI vesicles recycle Golgi resident proteins from later to earlier cisternae, and the nature of the cisternae gradually changes as they progress (maturation). This model was supported by the fact that the cargoes much larger than COPI vesicles can travel across the Golgi stacks. Cell surface scales of algae are assembled in the Golgi apparatus, and transported forward without being packed into COPI vesicles (Becker et al., 1995; Brown et al., 1970; Melkonian et al., 1991). Procollagen aggregates in mammalian fibroblasts are also shown to progress across the Golgi stacks without getting into small vesicles (Bonfanti et al., 1998; Mironov et al., 2001). One important problem here is the direction of intra-Golgi COPI vesicles. Many studies have been made using mammalian cells to solve this problem, but have led to conflicting conclusions. Some found that Golgi glycosylation enzymes were enriched in COPI vesicles but cargo proteins were not (Gilchrist et al., 2006; Martıńez-Mena´rguez et al., 2001), whereas others reported the opposite (Cosson et al., 2002; Kweon et al., 2004; Orci et al., 2000a). Recent studies on the tethering factors involved in intra-Golgi movement of COPI vesicles revealed that


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untethered COPI vesicles with Golgi resident proteins accumulated when they were knocked down (Sohda et al., 2010; Zolov and Lupashin, 2005). Only retrograde but not anterograde trafficking was disrupted in these experiments, indicating the involvement of COPI vesicles in the intra-Golgi retrograde trafficking. It was also suggested that there are at least two types of COPI vesicles according to their tethering factors, and this differentiation can be coupled with their transport directions (forward/bidirectional and backward) (Malsam et al., 2005; Orci et al., 2000b). However, the two COPI populations could be retrograde carriers, and the existence of anterograde vesicles remains to be confirmed. Recent data from S. cerevisiae by high-resolution confocal microscopy have revealed that the protein components change in a single cisterna from cis-Golgi proteins to trans-Golgi proteins over time, which gives a strong support for the maturation of cisternae (Losev et al., 2006; MatsuuraTokita et al., 2006). Detailed morphological studies by electron tomography of the cells of human, Pichia pastoris, Arabidopsis thaliana, Scherffelia dubia, Chlamidomonas reinhardtii, and Dionaea muscipula (Venus flytrap) support cisternal maturation (Donohoe et al., 2013; Mogelsvang et al., 2003; Storrie et al., 2012). Moreover, observations of Golgi regeneration after BFA treatment and removal in mammalian and plant cells have revealed that the Golgi stacks regenerate in the cis-to-trans order, which is consistent with the cisternal maturation model (see Section 5.4; Alcalde et al., 1992; Ito et al., 2012; Puri and Linstedt, 2003; Schoberer et al., 2010). Therefore, the basic concept of the cisternal maturation model appears to be widely accepted now. However, the debate has not come to an end yet. Patterson et al. (2008) reported that both big and small secretory cargoes exit the Golgi with exponential kinetics in mammalian cells. This appeared to conflict with the cisternal maturation, in which the cargoes are predicted to exit the Golgi with linear kinetics. The problem can be avoided by modifying the model to include a post-Golgi compartment such as TGN as longlived organelle, but so far there is no supporting evidence. Moreover, recent two studies using similar procedures report completely opposite results (Lavieu et al., 2013; Rizzo et al., 2013). The two groups used the domain which can be polymerized at will, and one group fused it with the CTS region of Golgi mannosidase and the other fused it with the luminal side of a plasma membrane protein. The polymerized protein of the former group progressed forward across the Golgi stack,

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whereas that of the latter group stayed and was not transported to later cisternae. The debate is still going on, and we obviously need further investigations.

4.2. Tethering and fusion of intra-Golgi COPI vesicles For tethering of intra-Golgi COPI vesicles, although the precise origin and transport direction of the vesicles are not clear, the coiled-coil tethers p115 (yeast Uso1), giantin, golgin-84, and CASP (yeast Coy1), and the multisubunit complexes TRAPPII and COG are suggested to be involved. Uso1 is essential only for COPII tethering as described in Section 3.2.3, but mammalian p115 additionally binds to b-COP and is involved in COPI tethering (Guo et al., 2008). Giantin and golgin-84 bind to p115 and CASP, respectively, on the Golgi membranes (Malsam et al., 2005). COG is an octameric complex composed of Cog1–8 subunits, and yeast Cog2 and human Cog3 have been reported to interact with g- and b-COP, respectively (Suvorova et al., 2002; Sztul and Lupashin, 2009; Vasile et al., 2006; Zolov and Lupashin, 2005). Yeast COG complex strongly interacts with active Ypt1, and also with Ypt6 to a less extent independently of nucleotides (Suvorova et al., 2002). Similarly, mammalian COG complex interacts with Rab1 and Rab6, the homologs of Ypt1 and Ypt6 (Fukuda et al., 2008). Not only p115 and COG complex but also giantin and golgin-84 interact with active Rab1 (or yeast Ypt1; Beard et al., 2005; Diao et al., 2003; Fukuda et al., 2008; Satoh et al., 2003). Mammalian TRAPPII is suggested to act not only as a tether but also as a specific GEF of Rab1 similarly to TRAPPI, and the activated Rab1 recruits effectors including the tethering factors above (Allan et al., 2000). TRAPPI and TRAPPII complexes share common subunits, and TRAPPII has additional three subunits, which are thought to determine the binding specificity different from TRAPPI (Sztul and Lupashin, 2009). Mammalian TRAPPII complex is especially associated with COPI vesicles and buds, and its subunits interact with g-COP (Yamasaki et al., 2009). In plants, homologs of p115, golgin-84, CASP, and COG complex components have been found, but not for giantin and the TRAPPII subunits that are not shared with TRAPPI (Latijnhouwers et al., 2005). Although the role of Atp115 in ER-to-Golgi anterograde transport is


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indicated as described in Section 3.2.3, it is unknown whether it mediates COPI tethering. Homologs of golgin-84 were identified as GC1 and GC2, and their GFP-fusion proteins localized to the cis-Golgi cisternal rims (Latijnhouwers et al., 2007). The Arabidopsis CASP homolog AtCASP was identified based on similarity to human CASP in transmembrane region, and localized to the Golgi cisternal rims in tobacco leaf epidermal cells (Latijnhouwers et al., 2007; Renna et al., 2005). This localization was later revealed to be the cisternae between those with the cis-Golgi SNARE AtSYP31 and the trans-Golgi marker ST in tobacco BY-2 cells (Ito et al., 2012). The physiological roles of GC1, GC2, and AtCASP in plant cells have not been reported. For COG complex, homologs of all subunits are coded in the Arabidopsis genome (Latijnhouwers et al., 2005). Among the eight subunits, the gene coding Cog7 has been shown to be responsible for embryo yellow (eye) mutant, which exhibits defects in cell expansion and organization of the shoot apical meristem. In eye mutant embryos, ERD2-GFP mislocalizes to the ER and GFP-tagged cis-Golgi localized SNARE MEMB12 shows aberrant shaped punctate structures (Ishikawa et al., 2008). In addition to Rab1 homolog RABDs (see Section 3.2.3), Rab6/Ypt6 group is also conserved in plants; the Arabidopsis homologs are called RABH group (Saito and Ueda, 2009). Because one of the RABH proteins AtRABH1b has been reported to replace the function of yeast Ypt6, the molecular function is thought to be conserved (Bednarek et al., 1994). After tethering, membrane fusion is thought to be mediated by SNARE proteins; Qa-SNARE Sed5 (mammalian syntaxin 5), Qb-SNARE Gos1 (Gos28), Qc-SNARE Sft1 (GS15), and R-SNARE Ykt6 or Sec22 (Barlowe and Miller, 2013; Kloepper et al., 2007; Shorter et al., 2002; Xu et al., 2002). Among the plant Golgi-localized SNAREs (see Section 3.2.3), AtSYP3, AtGOS1, and AtBS14 families, and AtSEC22 are presumed to be involved in intra-Golgi COPI tethering from their homology to yeast SNAREs (Sanderfoot et al., 2000; Tai and Banfield, 2001). Because overexpression of AtBS14a or At BS14b has been shown to partially suppress the phenotypes of yeast sft1 deletion mutant, their functions are thought to be conserved to some extent (Tai and Banfield, 2001). For AtSYP31 and AtSEC22, although their involvement in ER–Golgi trafficking is speculated, there is no experimental data indicating its role in intraGolgi transport.

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5. FORMATION AND MAINTENANCE OF STACKED STRUCTURE 5.1. Golgi apparatus as stacked cisternae As described in Section 2.1, the unique stacked structure of the Golgi apparatus is conserved among nearly all eukaryotes. Certainly, there are a few exceptions such as S. cerevisiae, which has unstacked Golgi, and some unicellular parasites, which appear to lack Golgi. Many of such “Golgi-lacking” organisms also lack identifiable mitochondria and peroxisomes, and they were thought to have evolved very early before the innovation of the Golgi in the early studies (Cavalier-Smith, 1987). However, improved phylogenetic analyses revealed later that all of them possess most genes encoding the proteins associated with trafficking around the Golgi such as coat proteins, Rab GTPases, and SNAREs. Moreover, the organisms without stacked Golgi like S. cerevisiae are spread throughout the eukaryotic tree, each of which is located in the clade of organisms with the Golgi stacks (Klute et al., 2011; Mowbrey and Dacks, 2009). Thus, the loss of the normal Golgi structures is now thought to be secondary events. The phylogenetic tree suggests that Golgi stacking has been lost at least eight times independently (Mowbrey and Dacks, 2009). Thus, the ancestral eukaryote must have had stacked Golgi and the structure has been inherited to most descendants during evolution. This indicates that the stacked structure is important for the Golgi function and therefore for better survival of eukaryotes. The relationship between the Golgi structure and functions has been asked for a long time. Cisternal stacking may contribute to efficient transport through the Golgi apparatus, because the donor and target membranes are physically close to each other. However, S. cerevisiae without stacked Golgi achieves efficient protein secretion (Nakano and Luini, 2010). Also in Drosophila, the cells at many developmental stages are known to exhibit no Golgi stacks but instead possess vesiculotubular clusters, still achieving efficient secretion (Kondylis and Rabouille, 2009). By contrast, Golgi stacking might limit the intra-Golgi transport. COPI vesicles are known to form exclusively at the cisternal rims, and different behaviors between the rims and the center of the cisternae are proposed (Lavieu et al., 2013). Golgi unstacking can increase the membrane surface of cisternae, which facilitates interaction with coat proteins to produce transport vesicles. In support of this, artificially unstacked mammalian


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Golgi was reported to produce vesicles more efficiently than normal Golgi stacks in vitro (Wang et al., 2008). From another viewpoint, Golgi stacking is presumed to ensure complex cargo glycosylation by sequential exposure of cargo to enzymes. Some “Golgi-lacking” organisms are suggested to possess fewer ER glycosyltransferases compared to their relatives with Golgi stacks, implying their difference in Golgi glycosylation as well (Samuelson et al., 2005). A recent study shows that disruption of the Golgi structure accelerates trafficking but impairs proper cargo glycosylation (Xiang et al., 2013). Further comparative studies will give us more insights.

5.2. Search for the “Golgi stacking factors” In order to form and maintain the stacked structure of the Golgi apparatus, there should be molecular factors responsible for it. Such components have been sought for a long time. By electron microscopy, bridge-like components cross-linking the adjacent cisternae were observed in animal and plant cells, which turned out to be proteins sensitive to proteases (Cluett and Brown, 1992; Franke et al., 1972; Mollenhauer, 1965; Staehelin et al., 1990). To identify these components, the procedures used for characterization of nuclear lamina were applied; membranes of isolated Golgi stacks were solubilized by detergent, and after removal of Golgi enzymes so-called Golgi matrix was isolated (Slusarewicz et al., 1994). Antisera raised against this matrix led to identification of GM130 (Nakamura et al., 1995). An in vitro assay isolated another component. Mammalian Golgi apparatus disassembles upon mitosis and reassembles in the daughter cells at telophase, and this Golgi disassembly and reassembly can be mimicked in a cell-free assay treating the isolated Golgi apparatus with mitotic or interphase cytosol. An alkylating agent NEM (N-ethylmaleimide) prevented Golgi reassembly, and its major target turned out to be a 65-kDa protein. The protein was named GRASP65 (Golgi ReAssembly Stacking Protein), because inhibiting its function by addition of its antibody or recombinant GRASP65 after Golgi disassembly led to the formation of single cisternae without stacking upon reassembly (Barr et al., 1997). GRASP65 was revealed to be enriched in the detergent-resistant matrix fraction and interact with GM130 (Barr et al., 1998). Identification of GRASP65 led to the discovery of a homolog GRASP55 with a similar role (Shorter et al., 1999). The in vivo function of GRASP65 in Golgi stacking was supported to some extent by the experiments of microinjection of the antibody against

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GRASP65 and gene knockdown by RNAi (Su¨tterlin et al., 2005; Wang et al., 2003). However, upon inhibition of either GRASP65 or GRASP55 expression by siRNA, the Golgi ribbons fragmented into ministacks but Golgi stacking remained unaffected (Feinstein and Linstedt, 2008; Puthenveedu et al., 2006). Subsequently, double depletion of GRASP65 and GRASP55 by siRNA was performed, and electron microscopy revealed that stacked Golgi was absent in over 80% of the cells (Xiang and Wang, 2010). From these results, mammalian GRASPs are now thought to act in Golgi stacking redundantly, but independently in Golgi ribbon formation. In both cases, GRASPs probably function as bridges between adjascent compartments by oligomerization (Lowe, 2011). GM130 is also suggested to be required for both Golgi stacking and Golgi ribbon formation, presumably as the tethering factor acting before connection by GRASPs (Marra et al., 2007; Puthenveedu et al., 2006; Shorter and Warren, 1999). In spite of their prominent roles in the Golgi structure formation in mammalian cells, the function of GRASP homologs as the “Golgi stacking protein” is not very obvious in other organisms. Unlike vertebrates with clear Golgi ribbons, most organisms with discrete Golgi stacks (or no stacks) possess only one GRASP homolog (Vinke et al., 2011). In Drosophila cells, depletion of the only GRASP homolog dGRASP effects only 30% of the Golgi stacks, whereas double depletion with GM130 homolog increases the effect (Kondylis et al., 2005). In addition, dGRASP localizes to the Golgi membranes even when the Golgi cisternae are unstacked (Kondylis and Rabouille, 2009). Deletion of the GRASP homolog Grh1 in P. pastoris has no obvious effect on the Golgi structure (Levi et al., 2010). S. cerevisiae also has a GRASP homolog, which localizes to the cis-Golgi membrane, in spite of the well-known unstacked Golgi (Behnia et al., 2007). Surprisingly, GRASP is lacking in plants. Green algae also lack GRASP but red algae have it, indicating the secondary loss of GRASP in the chloroplastida stem lineage (Klute et al., 2011). Similarly, GM130 has not been found in plants either (Latijnhouwers et al., 2007). Needless to say, however, plants have stacked Golgi apparatus. Whether conservation of the genes involved is too low or completely different components substitute GRASP and GM130 in plants remain obscure. The plant Golgi stacks keep intact structure during their rapid movement and never break down while traveling (Madison and Nebenfu¨hr, 2011). The best candidate that keeps the structural integrity is the Golgi matrix proteins. By electron microscopy, plant cells show a zone surrounding the Golgi


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stack and the TGN, from which ribosomes are excluded (Staehelin and Kang, 2008; Staehelin et al., 1990). This “ribosome-exclusion zone” is thought to represent the Golgi matrix holding the Golgi structure together. Some matrix components have been identified in plant cells, including AtCASP and Atp115, which also act as vesicle tethering factors (Sections 3.2.3 and 4.2), but their functions in the maintenance of the Golgi structure are not known. Further investigations would reveal the mechanism for Golgi structure maintenance in plant cells, perhaps not much conserved with animals and yeast.

5.3. Golgi biogenesis upon mitosis Upon cell division, not only DNA but also organelles must be inherited to the daughter cells. The Golgi apparatus is not an exception. Some parasitic protozoa have only one Golgi stack per cell, which makes it easy to track the behaviors of the single Golgi. By several studies, the Golgi apparatus in such organisms are shown to double before cytokinesis by distinct procedures. In Trichomonas, the Golgi stack elongates laterally during cell cycle and undergoes medial fission before mitosis. The fission appears to occur from trans-to-cis direction in electron microscopy (Benchimol et al., 2001). In Toxoplasma gondii, similar lateral growth and fission are followed by an additional fission. The resulting four stacks are partitioned to the daughter cells in pairs, which subsequently coalesce to form a single Golgi in each daughter cell (Pelletier et al., 2002). Differently from these species, the new Golgi stack is formed de novo near the old Golgi in Trypanosoma brucei. The Golgi formation is coupled with duplication of the ERES, and the new Golgi stack is supplied with materials not only by the new ERES but also by the old Golgi (He et al., 2004). The new Golgi formation was visualized by using several Golgi proteins, showing that the Golgi matrix and enzymes are laid down first, followed by the association of COPI components and initiation of cargo trafficking (Ho et al., 2006). Such de novo formation of the ERES– Golgi unit has been also observed in P. pastoris, whereas the Golgi formation is not limited around mitosis. This process is seemingly independent of preexisting Golgi stacks, but it is also possible that there are invisible Golgi templates (Bevis et al., 2002; Rossanese et al., 1999). In contrast to the protozoans, the mammalian Golgi apparatus behaves in a completely different way. The single Golgi ribbon is initially disassembled into the ministacks in the late G2 phase, and even the stacks disappear at the early phase of mitosis. What occurs in the latter step is still under debate;

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whether the Golgi cisternae fragment into small vesicles, or the Golgi components are absorbed into the ER (Axelsson and Warren, 2004; Persico et al., 2009; Zaal et al., 1999). In either case, the Golgi components are dispersed throughout the cell and partitioned into the daughter cells, followed by reassembly of the Golgi ribbon in the telophase. The reassembly process has been intensively studied to reveal the mechanism of Golgi biogenesis, and led to the identification of GRASPs as described in the previous section. In plant cells, it is known that the Golgi stacks remain intact all through the cell cycle (Dupree and Sherrier, 1998; Nebenfu¨hr et al., 2000). Whether the Golgi apparatus disassembles or remains intact during mitosis is not species dependent but cell-type dependent, because Drosophila Golgi undergoes disassembly and reassembly in tissue cultured cells but keeps intact in embryonic cells (Stanley et al., 1997). It is hypothesized that the difference might depend on the ER–Golgi transport activity. During mitosis, Sec13 dissociates from the ERES and the ER export is arrested in mammalian cells (Hammond and Glick, 2000). On the other hand, plant cells require active secretion for cytokinesis, which is achieved by the cell plate. Cell plate formation needs new membrane and wall materials supplied by Golgi-derived vesicles to the phragmoplast, therefore disruption of the Golgi apparatus by BFA during mitosis results in binuclear cells (Reichardt et al., 2007). Because the plant Golgi stacks do not disassemble at any stages, they must somehow increase in number during the cell cycle. However, the process of Golgi biogenesis and its relationship to mitosis are almost completely unknown in plant cells. In onion root meristem cells, there is a report that the Golgi number increases mainly between prophase and metaphase (Garcia-Herdugo et al., 1988). On the other hand, the Golgi stacks in Catharanthus roseus cells laterally enlarge during cytokinses, and some stacks seemed to divide by electron microscopy (Hirose and Komamine, 1989). By electron tomography, the “dividing” Golgi looks as two stacks sharing a big trans-cisterna (Staehelin and Kang, 2008). According to the cisternal maturation model, such division should start with assembly of two half-sized cis-cisternae on the cis-surface of single enlarged stack, and maturation of the cis-cisternae would progress without fusion being followed by assembly of new cisternae, each of which eventually results in two separated stacks. However, electron microscopy cannot distinguish whether the stacks are dividing or fusing. A careful stereological study by electron microscopy in Arabidopsis shoot apical meristem cells has suggested that the Golgi stacks doubles in number during the G2 phase prior to mitosis, but its mechanism


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remains unknown (Seguı´-Simarro and Staehelin, 2006). Live cell imaging analysis also has been performed, however, no dramatic increase of the number of the Golgi has been observed in any specific time period (Nebenfu¨hr et al., 2000).

5.4. Golgi regeneration after BFA treatment As described above, BFA treatment causes disassembly of the Golgi apparatus and absorbance of most Golgi components into the ER in many types of cells, and this effect is reversible; the Golgi apparatus reassembles after BFA removal. This reassembly process has been investigated to reveal the mechanism of Golgi biogenesis. Especially in mammalian cells, many studies using BFA have been performed to know whether the Golgi biogenesis occurs de novo from the ER or requires preexisting templates. The candidate of such templates is the Golgi matrix, because it is segregated from other Golgi components including Golgi enzymes by BFA treatment, localizing to the “Golgi remnants” distinctive from the ER. In addition, mitosis progresses normally under this condition, and the Golgi apparatus is regenerated by BFA removal after cell division (Seemann et al., 2002). However, combination of BFA and another drug H89, which blocks ER export, induces ER redistribution of both matrix and nonmatrix proteins, and the Golgi reassembles normally by removing them (Puri and Linstedt, 2003). Therefore, the mammalian Golgi is thought to have the ability to form de novo under a certain condition. In plant cells, the behavior of the Golgi proteins upon BFA treatment has been controversial. In tobacco leaf epidermal cells, the Golgi proteins were reported to redistribute to the ER in the trans-to-cis order by BFA treatment. In addition, one of the matrix protein, AtCASP, did not fully relocate to the ER but localized to small punctate structures colocalizing with ERES (Schoberer et al., 2010). On the other hand, we and other have shown that both cis- and trans-Golgi cisternae collapse almost at the same time in tobacco BY-2 cultured cells (Ito et al., 2012; Madison and Nebenfu¨hr, 2011). Moreover, we have demonstrated that AtCASP relocalizes to the ER almost completely by BFA treatment (Ito et al., 2012). We have also demonstrated that the cis-Golgi proteins AtSYP31 and AtRER1B relocalize to punctate structures by BFA treatment in BY-2 cells, whereas another cis-Golgi protein AtERD2 is absorbed into the ER (Ito et al., 2012). Very similar results were obtained in Arabidopsis cultured cells by transient expression of dominant mutants of ARF1 (Takeuchi et al., 2002).

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The punctate structures of AtSYP31 and AtRER1B are smaller in size and larger in number compared to normal Golgi stacks, and detailed 3D analysis by super-resolution confocal live imaging microscopy (SCLIM) that we had developed has revealed that these structures lie very close to the ERES but are not identical to the ERES (Ito et al., 2012). Relation of these punctate structures with those of AtCASP observed in tobacco leaf epidermal cells (Schoberer et al., 2010) is unknown. It is also reported that expression of the GTP-locked dominant mutant of SAR1 in tobacco cells results in Golgi fragmentation and accumulation of vesiclulotubular clusters as the remnants (Osterrieder et al., 2010). Such clusters might be related to the punctate structures we observe, but further investigations are needed. After BFA removal, the punctate structures of the cis-Golgi proteins gather first, and subsequently the medial- and trans-Golgi markers follow sequentially to form Golgi stacks (Ito et al., 2012). This result indicates that the punctate structure function as “seeds” of Golgi regeneration by receiving cis-Golgi materials to form the cis-cisternae, and regeneration progresses in the cis-to-trans order (Fig. 6.7). According to a report by electron tomography, the cis-most cisternae in plant cells are biosynthetically inactive and might function similarly to mammalian ERGIC (Donohoe et al., 2013). In mammalian cells upon BFA treatment, not only matrix proteins but also some cis-Golgi membrane proteins that cycle between the ER and the Golgi and also the ERGIC protein ERGIC53 localize to the Golgi remnants (Fu¨llekrug et al., 1997a,b; Lippincott-Schwartz et al., 1990; Tang et al., 1993). From these facts, we speculate that the cis-most cisternae in plant cells function like both the ERGIC and the cis-Golgi of mammalian cells, and the proteins localizing there relocate to the punctate structures by BFA treatment that act as the scaffold for Golgi regeneration. In the later stage of Golgi regeneration after BFA removal, the Golgi stacks keep growing until they reach the size twice as large as normal Golgi, and subsequently divide by fission (Hawes et al., 2008a; Langhans et al., 2007). By electron microscopy, the fission seems to occur in the cis-to-trans order, consistent with the fission by maturation (Section 5.3). It is still unknown in which way the new Golgi is formed (duplication by fission, de novo formation from the ER, or the intermediate), but these results from the Golgi regeneration experiments indicate that any of them can occur in plant cells, although the final division has not been observed in living cells. Further studies by live imaging with higher time and space resolution will provide us more insights.


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Figure 6.7 A model for the behaviors of the tobacco Golgi apparatus upon BFA treatment and removal. (A) The Golgi stacks are located near the ERES, and there might be some ERES without associated Golgi. (B) By BFA treatment, some cis-Golgi proteins localize to the punctate compartments closely associated to the ERES, while other cis-, medial-, and trans-Golgi proteins relocate to the ER membrane. (C) By BFA removal, the punctate compartments with cis-Golgi proteins gather first to form the cis-cisternae. The Golgi stacksreassemble from the regenerated cis-cisternae in the cis-to-trans order (back to A).

6. RELATIONSHIP BETWEEN THE GOLGI APPARATUS AND THE TGN The TGN was first defined in mammalian cells as a specialized compartment on the trans-side of the Golgi apparatus composed of tubular networks (Griffiths and Simons, 1986; Roth et al., 1985; Taatjes and Roth, 1986). Similarly to the Golgi cisternae, the TGN also contains many resident enzymes for cargo processing (Rabouille et al., 1995). However, in addition to function as the sequel to the cargo processing line of the Golgi, the TGN sorts the cargo proteins destined for the plasma membrane and endosomes (Keller and Simons, 1997). Moreover, the TGN receives cargo from endosomal compartments (Pavelka et al., 1998). Thus, the TGN acts as the trafficking hub that both the secretory and endocytic pathways pass through. The Golgi cisternae and the TGN can be distinguished by the vesicles they form, because the Golgi cisternae generate COPI vesicles

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but not clathrin-coated vesicles, and the opposite is true for the TGN (Ladinsky et al., 2002). In plant cells, the TGN was first identified as “partially coated reticulum,” a vesiculotubular compartment with clathrin-coated buds (Pesacreta and Lucas, 1985), and later the word TGN was applied from mammalian studies (Staehelin et al., 1990). Similarly to the mammalian TGN, clathrin-coated vesicles are formed from the TGN, but not from the Golgi cisternae in plant cells (Donohoe et al., 2007). The plant TGN is marked by several SNARE proteins including SYP41, SYP61, and VPS45 (Uemura and Nakano, 2013). Using SYP41 and a subunit of V-ATPase (VHAa1) as the TGN markers, it has been demonstrated that a endocytosis tracer FM4–64 localizes to the TGN before late endosomes, indicating that the TGN functions as an early endosome (Dettmer et al., 2006). This is also supported by the observations by time-lapse imaging and immunoelectron microscopy, which have shown that the plasma membrane proteins internalized by endocytosis pass through the TGN (Viotti et al., 2010). In addition to the functional difference, dynamic behavior of the TGN is distinct from the Golgi cisternae. By electron tomography in Arabidopsis cells, the TGN seems to get separated from the Golgi (Staehelin and Kang, 2008). Such “free-floating TGN” is also reported in P. pastoris (Mogelsvang et al., 2003). Further observation by spinning disc confocal microscopy has revealed that the plant TGN moves independently of the Golgi apparatus and transiently associates with it (Viotti et al., 2010). Our detailed analysis by SCLIM has also demonstrated that there are two types of TGN; one is associated with the Golgi stacks and the other is independent from the Golgi (Uemura et al., in press). Such differences between the Golgi apparatus and the TGN may reflect their protein composition. Now it is possible to isolate and characterize the Golgi and TGN proteins separately. A recent study suggests that almost 30% of the proteins identified in the Arabidopsis TGN are non-Golgi proteins (Parsons et al., 2013). It is also known that the Golgi and the TGN can be distinguished by different behaviors upon BFA treatment. The formation of clathrin-coated vesicles at the TGN is thought to be regulated by ARF1, whose GEFs are affected by BFA. In mammalian cells, although the Golgi components are absorbed into the ER by BFA treatment, the TGN fuses with endosomes (Lippincott-Schwartz et al., 1991; Wood et al., 1991). In Arabidopsis root cells, in which the Golgi stacks do not disappear by BFA, the TGN forms large aggregates called BFA bodies (or BFA compartments) with endosomal


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components (Langhans et al., 2011; Uemura and Nakano, 2013). On the other hand, in tobacco cells, in which the Golgi stacks fuse and get absorbed into the ER upon BFA treatment, the TGN forms what look like BFA bodies, which are much smaller than those in Arabidopsis root cells (Langhans et al., 2011). This difference may be because the Golgi maturation continues and supplies materials to the TGN in the presence of BFA in Arabidopsis root cells but not in tobacco cells. Our detailed time-lapse observation tobacco BY-2 cells has also revealed that the regeneration process of the TGN after BFA removal is largely independent from the Golgi apparatus, but the association with the Golgi is indispensable (Ito et al., unpublished). These results indicate the complex nature of the plant TGN functioning both in the secretory and endocytic pathways.

7. CONCLUDING REMARKS As the application of electron microscopy promoted the Golgi apparatus from an artifact to the center of attention in cell biology, the improvement of light microscopy is bringing us the new perspective about the organelle being highly dynamic. Especially in these years, we are overcoming the diffraction limit of light microscopy by super-resolution microscopes. The plant Golgi apparatus provides an ideal system to apply such techniques, which will give us new insights about the structural organization and trafficking mechanisms of the Golgi stacks. Moreover, in spite of the high conservation of the stacked structure, the molecular components accomplishing the Golgi stacking appear to be different from organism to organism. The physiological significance of the rapid movement of the plant Golgi stacks throughout the cell is still mysterious. Further investigation of the plant Golgi apparatus including analysis of their cargo will shed light to the Golgi functions unique and important for plant life.

ACKNOWLEDGMENTS We thank members of the Nakano laboratory for valuable discussions. This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science, and Technology of Japan and by funds from the Extreme Photonics and Cellular Systems Biology Projects of RIKEN. Y.I. is a recipient of the Research Fellowship for Young Scientists from the Japan Society for the Promotion of Science.

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Specific organization of Golgi apparatus in plant cells.

Unique Golgi apparatus and vesicle formation in a red alga.

Formation of secretory granules in the Golgi apparatus of prolactin cells in the rat pituitary gland: a stereoscopic study.

Separation of the plant Golgi apparatus and endoplasmic reticulum by free-flow electrophoresis.

The centrosome-Golgi apparatus nexus.

Formation of wheat protein bodies: Involvement of the Golgi apparatus in gliadin transport.

Isolation of a Golgi-apparatus-enriched fraction from leukaemic cells.

Cytochemical characteristics of the Golgi apparatus.

Role of microtubules in the organisation of the Golgi apparatus.

Three-dimensional architecture of the golgi apparatus in Sertoli cells of the rat.

Intracellular zinc distribution in mitochondria, ER and the Golgi apparatus.

Perspectives on Golgi apparatus form and function.

Role of the Golgi apparatus in cellular pathology.

Mannosylation of fungal glycoconjugates in the Golgi apparatus.

Golgi apparatus of epithelial principal cells of the epididymal initial segment of the rat: structure, relationship with endoplasmic reticulum, and role in the formation of secretory vesicles.

Newly synthesized acetylcholine receptors are located in the Golgi apparatus.

The lipids of the Golgi apparatus subfractions from rat liver.

Cell biology: Polarized transport in the Golgi apparatus.

Endoplasmic reticulum-Golgi apparatus relationships in the rat spermatid.

Golgi apparatus and protein trafficking in Alzheimer's disease.

The Golgi apparatus of rat pachytene spermatocytes during spermatogenesis.

Growth of the Mammalian Golgi Apparatus during Interphase.

Traffic through the Golgi apparatus as studied by radioautography.

Mad1 at the Golgi apparatus: a story beyond kinetochores.