Microtubules Contribute to Tubule Elongation and Anchoring of Endoplasmic Reticulum,Resulting in High Network Complexity in Arabidopsis

The endoplasmic reticulum (ER) is a network of tubules and sheet-like structures in eukaryotic cells. Some ER tubules dynamically change their morphology, and others form stable structures. In plants, it has been thought that the ER tubule extension is driven by the actin-myosin machinery. Here, we show that microtubules also contribute to the ER tubule extension with an almost 20-fold slower rate than the actin filament-based ER extension. Treatment with the actin-depolymerizing drug Latrunculin B made it possible to visualize the slow extension of the ER tubules in transgenic Arabidopsis (Arabidopsis thaliana) plants expressing ER-targeted green fluorescent protein. The ER tubules elongated along microtubules in both directions of microtubules, which have a distinct polarity. This feature is similar to the kinesin- or dynein-driven ER tubule extension in animal cells. In contrast to the animal case, ER tubules elongating with the growing microtubule ends were not observed in Arabidopsis. We also found the spots where microtubules are stably colocalized with the ER subdomains during long observations of 1,040 s, suggesting that cortical microtubules contribute to provide ER anchoring points. The anchoring points acted as the branching points of the ER tubules, resulting in the formation of multiway junctions. The density of the ER tubule junction positively correlated with the microtubule density in both elongating cells and mature cells of leaf epidermis, showing the requirement of microtubules for formation of the complex ER network. Taken together, our findings show that plants use microtubules for ER anchoring and ER tubule extension, which establish fine network structures of the ER within the cell.The endoplasmic reticulum (ER) is a complex network composed of tubules and sheet structures. The ER network’s morphology changes dynamically by elongation and shrinkage of tubules, sheet expansion, and sliding junctions. For example, an ER tubule elongates straight forward from a cisterna and subsequently, fuses to another cisterna, producing a linkage between two cisternae. If an elongating tubule fails to fuse to another cisterna, the tubule contracts into the original cisterna. However, the ER has stable anchoring points that associate with other cellular structures, such as the plasma membrane or cytoskeleton. When an elongating ER tubule reaches an association point, it forms a stable ER anchor (i.e. establishment of the ER anchoring points forms stable ER tubules). Hence, increasing the number of ER anchoring points produces fine ER meshwork.ER dynamics in eukaryotes depend on the cytoskeleton. In plants, major contributors for ER organization are actin filaments (Quader et al., 1989; Knebel et al., 1990; Lichtscheidl and Hepler, 1996; Sparkes et al., 2009a) and the actin-associated motor proteins (myosins; Prokhnevsky et al., 2008; Peremyslov et al., 2010; Ueda et al., 2010). However, it had generally been thought that microtubules are not involved in ER organization in plants, because microtubule-depolymerizing drugs do not induce obvious changes in the ER network (Quader et al., 1989; Knebel et al., 1990; Lichtscheidl and Hepler, 1996; Sparkes et al., 2009a). Nevertheless, involvement of microtubules in plant ER organization has been suspected from several electron microscopy observations that showed microtubules located close to the ER membrane in Vicia faba guard cells, Nicotiana alata pollen tubes, and Funaria hygrometrica caulonemata (Lancelle et al., 1987; Hepler et al., 1990; McCauley and Hepler, 1992).Foissner et al. (2009) have suggested that microtubules are involved in motility and orientation of cortical ER in Characean algae (Nitella translucens, Nitella flexilis, Nitella hyalina, and Nitella pseudoflabellata) internodal cells. Characean cortical ER is spatially separated from inner cytoplasmic streaming by the middle layer of fixed chloroplasts. The cortical ER forms a tight meshwork of predominantly transverse ER tubules that frequently coalign with microtubules, and microtubule depolymerization reduces the transverse ER tubules and increases mesh size (Foissner et al., 2009). Consistently, Hamada et al. (2012) have shown in Arabidopsis (Arabidopsis thaliana) that microtubule depolymerization increases mesh size in young elongating cells. In addition, stable ER tubule junctions are often colocalized with cortical microtubules (Hamada et al., 2012), suggesting that microtubules stabilize ER tubule junctions to form fine ER meshes. Oryzalin-induced ER nodulation (Langhans et al., 2009) was not observed in our experimental conditions.Here, we showed that ER tubules elongate along microtubules in plant cells. In addition, we revealed that the ER is stably anchored to defined points on cortical microtubules. The stable anchoring points are the basis of various ER shapes, such as three-way, two-way, or dead-end ER tubules. These microtubule-ER interactions, together with the actin-myosin system, contribute to ER network organization.     

Reticulomics: Protein-Protein Interaction Studies with Two Plasmodesmata-Localized Reticulon Family Proteins Identify Binding Partners Enriched at Plasmodesmata,Endoplasmic Reticulum,and the Plasma Membrane

The endoplasmic reticulum (ER) is a ubiquitous organelle that plays roles in secretory protein production, folding, quality control, and lipid biosynthesis. The cortical ER in plants is pleomorphic and structured as a tubular network capable of morphing into flat cisternae, mainly at three-way junctions, and back to tubules. Plant reticulon family proteins (RTNLB) tubulate the ER by dimerization and oligomerization, creating localized ER membrane tensions that result in membrane curvature. Some RTNLB ER-shaping proteins are present in the plasmodesmata (PD) proteome and may contribute to the formation of the desmotubule, the axial ER-derived structure that traverses primary PD. Here, we investigate the binding partners of two PD-resident reticulon proteins, RTNLB3 and RTNLB6, that are located in primary PD at cytokinesis in tobacco (Nicotiana tabacum). Coimmunoprecipitation of green fluorescent protein-tagged RTNLB3 and RTNLB6 followed by mass spectrometry detected a high percentage of known PD-localized proteins as well as plasma membrane proteins with putative membrane-anchoring roles. Förster resonance energy transfer by fluorescence lifetime imaging microscopy assays revealed a highly significant interaction of the detected PD proteins with the bait RTNLB proteins. Our data suggest that RTNLB proteins, in addition to a role in ER modeling, may play important roles in linking the cortical ER to the plasma membrane.The endoplasmic reticulum (ER) is a multifunctional organelle (Hawes et al., 2015) and is the site of secretory protein production, folding, and quality control (Brandizzi et al., 2003) and lipid biosynthesis (Wallis and Browse, 2010), but it is also involved in many other aspects of day-to-day plant life, including auxin regulation (Friml and Jones, 2010) and oil and protein body formation (Huang, 1996; Herman, 2008). The cortical ER network displays a remarkable polygonal arrangement of motile tubules that are capable of morphing into small cisternae, mainly at the three-way junctions of the ER network (Sparkes et al., 2009). The cortical ER network of plants has been shown to play multiple roles in protein trafficking (Palade, 1975; Vitale and Denecke, 1999) and pathogen responses (for review, see Pattison and Amtmann, 2009; Beck et al., 2012).In plants, the protein family of reticulons (RTNLBs) contributes significantly to tubulation of the ER (Tolley et al., 2008, 2010; Chen et al., 2012). RTNLBs are integral ER membrane proteins that feature a C-terminal reticulon homology domain (RHD) that contains two major hydrophobic regions. These regions form two V-shaped transmembrane wedges joined together via a cytosolic loop, with the C and N termini of the protein facing the cytosol. RTNLBs can dimerize or oligomerize, creating localized tensions in the ER membrane, inducing varying degrees of membrane curvature (Sparkes et al., 2010). Hence, RTNLBs are considered to be essential in maintaining the tubular ER network.The ability of RTNLBs to constrict membranes is of interest in the context of cell plate development and the formation of primary plasmodesmata (PD; Knox et al., 2015). PD formation involves extensive remodeling of the cortical ER into tightly furled tubules to form the desmotubules, axial structures that run through the PD pore (Overall and Blackman, 1996; Ehlers and Kollmann, 2001). At only 15 nm in diameter, the desmotubule is one of the most constricted membrane structures found in nature, with no animal counterparts (Tilsner et al., 2011). PD are membrane-rich structures characterized by a close association of the plasma membrane (PM) with the ER. The forces that model the ER into desmotubules, however, are poorly understood. RTNLBs are excellent candidates for this process and can constrict fluorescent protein-labeled ER membranes into extremely fine tubules (Sparkes et al., 2010). We showed recently that two of the RTNLBs present in the PD proteome, RTNLB3 and RTNLB6 (Fernandez-Calvino et al., 2011), are present in primary PD at cytokinesis (Knox et al., 2015). However, nothing is known of the proteins that interact with RTNLBs identified in the PD proteome or that may link RTNLBs to the PM. To date, the only protein shown to bind to plant RTNLBs is RHD3-LIKE2, the plant homolog of the ER tubule fusion protein ATLASTIN (Lee et al., 2013).Here, we used a dual approach to identify interacting partners of RTNLB3 and RTNLB6 (Fernandez-Calvino et al., 2011; Knox et al.., 2015). First, we used GFP immunoprecipitation assays coupled to mass spectrometry (MS) to identify proteins potentially binding to RTNLB3 and RTNLB6. Second, from the proteins we identified, we conducted a detailed Förster resonance energy transfer by fluorescence lifetime imaging microscopy (FRET-FLIM) analysis to confirm prey-bait interactions in vivo.The application of time-resolved fluorescence spectroscopy to imaging biological systems has allowed the design and implementation of fluorescence lifetime imaging microscopy (FLIM). The technique allows measuring and determining the space map of picosecond fluorescence decay at each pixel of the image through confocal single and multiphoton excitation. The general fluorescence or Förster resonance energy transfer (FRET) to determine the colocalization of two color chromophores can now be improved to determine physical interactions using FRET-FLIM and protein pairs tagged with appropriate GFP fluorophores and monomeric red fluorescent protein (mRFP). FRET-FLIM measures the reduction in the excited-state lifetime of GFP (donor) fluorescence in the presence of an acceptor fluorophore (e.g. mRFP) that is independent of the problems associated with steady-state intensity measurements. The observation of such a reduction is an indication that the two proteins are within a distance of 1 to 10 nm, thus indicating a direct physical interaction between the two protein fusions (Osterrieder et al., 2009; Sparkes et al., 2010; Schoberer and Botchway, 2014). It was shown previously that a reduction of as little as approximately 200 ps in the excited-state lifetime of the GFP-labeled protein represents quenching through a protein-protein interaction (Stubbs et al., 2005).Our interaction data identified a large percentage (40%) of ER proteins, including other RTNLB family members. However, we also found a relatively large number (25%) of proteins present in the published PD proteome (Fernandez-Calvino et al., 2011) and a surprisingly high proportion (35%) of PM proteins. Of the PD-resident proteins we identified, a significant number were shown previously to be targets of viral movement proteins (MPs) or proteins present within lipid rafts, consistent with the view that PD are lipid-rich microdomains (Bayer et al., 2014). Additional proteins identified suggested roles for RTNLBs in transport and pathogen defense. We suggest that RTNLBs may play key roles in anchoring and/or signaling between the cortical ER and PM.     

Golgi- and Trans-Golgi Network-Mediated Vesicle Trafficking Is Required for Wax Secretion from Epidermal Cells

Lipid secretion from epidermal cells to the plant surface is essential to create the protective plant cuticle. Cuticular waxes are unusual secretory products, consisting of a variety of highly hydrophobic compounds including saturated very-long-chain alkanes, ketones, and alcohols. These compounds are synthesized in the endoplasmic reticulum (ER) but must be trafficked to the plasma membrane for export by ATP-binding cassette transporters. To test the hypothesis that wax components are trafficked via the endomembrane system and packaged in Golgi-derived secretory vesicles, Arabidopsis (Arabidopsis thaliana) stem wax secretion was assayed in a series of vesicle-trafficking mutants, including gnom like1-1 (gnl1-1), transport particle protein subunit120-4, and echidna (ech). Wax secretion was dependent upon GNL1 and ECH. Independent of secretion phenotypes, mutants with altered ER morphology also had decreased wax biosynthesis phenotypes, implying that the biosynthetic capacity of the ER is closely related to its structure. These results provide genetic evidence that wax export requires GNL1- and ECH-dependent endomembrane vesicle trafficking to deliver cargo to plasma membrane-localized ATP-binding cassette transporters.The aerial, nonwoody tissues of all land plants are covered by a waxy cuticle that protects the plant against nonstomatal water loss. The cuticle also provides the first barrier between the plant and its environment and mediates important biotic and abiotic interactions. The cuticle has two main components: cutin and waxes. Cutin is a tough, cross-linked polyester matrix primarily composed of C16 and C18 oxygenated fatty acids and glycerol (Pollard et al., 2008). Wax is a heterogenous mixture, primarily composed of very-long-chain (VLC) fatty acid derivatives (predominantly 29-carbon alkane in Arabidopsis [Arabidopsis thaliana] stems).As a result of biochemical approaches, forward genetic screens yielding the eceriferum (cer) mutants (Koornneef et al., 1989), and reverse genetics approaches (Greer et al., 2007), almost all of the enzymes in the wax biosynthesis pathway have been identified. The enzymes that elongate C16 or C18 fatty acids to VLC (greater than 20C) fatty acids are localized in the endoplasmic reticulum (ER; for review, see Haslam and Kunst, 2013). Primary alcohols are synthesized by the fatty acyl reductases (Rowland et al., 2006), while alkanes are generated via an undefined mechanism involving CER1, CER3, and an unidentified cytochrome b₅ (Bernard et al., 2012). These alkanes may be modified by the midchain alkane hydroxylase cytochrome P450 (MAH1) to generate secondary alcohols and ketones (Greer et al., 2007). All of these wax synthesis enzymes have also been localized to the ER (Greer et al., 2007; Bernard et al., 2012).In contrast to wax synthesis, comparatively little is known about how waxes are trafficked within the cell from their site of synthesis at the ER to the plasma membrane. ATP-binding cassette (ABC) transporters of the G subfamily are required for wax export, and when either half-transporter is disrupted, waxes accumulate in the ER (McFarlane et al., 2010). Two extracellular glycosylphosphatidylinositol-anchored lipid transfer proteins (LTPs) are further required for wax accumulation on the plant surface (DeBono et al., 2009; Kim et al., 2012). Although these components of the molecular machinery of wax transport at the plasma membrane have been identified, the intracellular mechanisms by which waxes are transported to the plasma membrane remain undefined.Several mechanisms have been hypothesized for the transport of waxes from the ER to the plasma membrane (for review, see Samuels et al., 2008). Waxes could be incorporated into vesicles at the ER, travel to and through the Golgi apparatus and the trans-Golgi network (TGN), and then move to the plasma membrane via vesicle secretion. These vesicles could carry wax components within their membranes, as computational modeling of wax components in lipid bilayers indicates that VLC alkanes partition entirely into the hydrophobic phase of the bilayer (Coll et al., 2007). Alternatively, lipoproteins may bind to lipid molecules in order to solubilize them so that they can be transported as cargo in the vesicle lumen, by analogy to mammalian systems where lipoproteins are secreted from hepatocytes into the circulatory system by exocytosis via post-Golgi vesicles (for review, see Mansbach and Siddiqi, 2010). However, no analogous lipid-binding apoproteins or transport vesicles have been found in plants. It is also possible that LTPs in membrane contact sites between the ER and the plasma membrane could transfer cuticular lipids directly from the ER to the plasma membrane. However, although these membrane contact sites have been observed in plant cells (Samuels and McFarlane, 2012), no structural or functional components of membrane contact sites are known.Early studies of VLC fatty acid trafficking used pulse-chase labeling to show that treatment with monensin, a post-Golgi trafficking inhibitor, results in decreased VLC fatty acid trafficking to the plasma membrane and a corresponding increase in these lipids in the Golgi apparatus (Bertho et al., 1991), suggesting a Golgi-dependent mechanism of VLC lipid trafficking to the plasma membrane. However, the “Golgi” fraction in this study contained significant elongase activity, which has subsequently been localized to the ER, making interpretation of these data difficult. While a variety of inhibitors are available that disrupt different stages in the secretory pathway (Zhang et al., 1993; Robinson et al., 2008), inhibitor studies of wax trafficking have proven ineffective, since the wax-producing epidermal cells do not effectively take up solutions carrying these inhibitors. This illustrates the difficulties of studying the transport of highly hydrophobic cargo, such as wax, within the single cell layer of epidermis.The objective of this study was to determine the intracellular trafficking mechanisms underlying cuticular wax transport from the ER to the plasma membrane. Arabidopsis mutants, which have been successfully applied in wax biosynthesis studies, were used to investigate wax secretion. Well-characterized mutants with defects in vesicle traffic and protein secretion were chosen to test the hypothesis that wax components are trafficked via endomembrane vesicles. These mutant analyses indicate that wax movement from the ER to the plasma membrane requires vesicle traffic at both the ER-Golgi interface and the TGN. Independent of secretion phenotypes, strong decreases in wax synthesis were observed in mutants with altered ER morphology, which implies that ER structure influences its biosynthetic capacity for wax production.     

Origin of β-Carotene-Rich Plastoglobuli in Dunaliella bardawil

The halotolerant microalgae Dunaliella bardawil accumulates under nitrogen deprivation two types of lipid droplets: plastoglobuli rich in β-carotene (βC-plastoglobuli) and cytoplasmatic lipid droplets (CLDs). We describe the isolation, composition, and origin of these lipid droplets. Plastoglobuli contain β-carotene, phytoene, and galactolipids missing in CLDs. The two preparations contain different lipid-associated proteins: major lipid droplet protein in CLD and the Prorich carotene globule protein in βC-plastoglobuli. The compositions of triglyceride (TAG) molecular species, total fatty acids, and sn-1+3 and sn-2 positions in the two lipid pools are similar, except for a small increase in palmitic acid in plastoglobuli, suggesting a common origin. The formation of CLD TAG precedes that of βC-plastoglobuli, reaching a maximum after 48 h of nitrogen deprivation and then decreasing. Palmitic acid incorporation kinetics indicated that, at early stages of nitrogen deprivation, CLD TAG is synthesized mostly from newly formed fatty acids, whereas in βC-plastoglobuli, a large part of TAG is produced from fatty acids of preformed membrane lipids. Electron microscopic analyses revealed that CLDs adhere to chloroplast envelope membranes concomitant with appearance of small βC-plastoglobuli within the chloroplast. Based on these results, we propose that CLDs in D. bardawil are produced in the endoplasmatic reticulum, whereas βC-plastoglobuli are made, in part, from hydrolysis of chloroplast membrane lipids and in part, by a continual transfer of TAG or fatty acids derived from CLD.Eukaryotic cells accumulate neutral lipids in different tissues mainly in the form of lipid droplets (Murphy, 2012). Most lipid droplets consist of a core of triglycerides (TAGs) and/or sterol esters coated by a phospholipids monolayer and embedded with proteins (Zweytick et al., 2000). Plants accumulate TAGs in different tissues, primarily in seeds but also in fruit, such as palm oil, flowers, and leaves. The best characterized system for TAG metabolism is oil seeds, in which TAG serves as the major carbon and energy reservoir to be used during germination (Huang, 1992, 1996). Recent studies show that lipid droplets are not just static pools of lipids but have diverse metabolic functions (Farese and Walther, 2009). In addition, plants also contain plastoglobuli, small chloroplastic lipid droplets consisting primarily of storage lipids and pigments. Proteome analyses of plastoglobuli suggest that they are involved in synthesis and degradation of lipids, pigments, and coenzymes (Ytterberg et al., 2006; Lundquist et al., 2012). It has been shown that plant plastoglobuli are associated with thylakoid membranes (Austin et al., 2006; Ytterberg et al., 2006).It is not entirely clear where the TAGs are synthesized in the plant cell. Until recently, it has been assumed that most TAGs are made in the endoplasmatic reticulum (ER) from fatty acids, which are mostly synthesized in the chloroplast and imported to the cytoplasm (Joyard et al., 2010). However, the recent identification of the enzyme diacylglycerol acyl transferase in plant plastoglobuli (Lundquist et al., 2012) suggests that TAG may be synthesized directly in chloroplasts, although direct evidence is missing. TAG may be synthesized also from galactolipid fatty acids during stress or senescence by phytyl ester synthases, which catalyze acyl transesterification from galactolipids to TAGs (Lippold et al., 2012). Phosphatidyl choline (PC) plays a major role in acyl transfer of newly synthesized fatty acids from the chloroplast into TAGs at the ER in plants (Bates et al., 2009). An indication for the origin of glycerolipids in plants is the identity of the fatty acids at the sn-2 position: if it originates in the chloroplast, it is mostly C16:0, whereas if it was made in the ER, it is mostly C:18 (Heinz and Roughan, 1983).Many species of unicellular microalgae can accumulate large amounts of TAGs under growth-limiting conditions, such as nitrogen deprivation (Shifrin and Chisholm, 1981; Roessler, 1990; Avron and Ben-Amotz, 1992; Thompson, 1996). In green microalgae (Chlorophyceae), TAGs are usually synthesized and accumulated in cytoplasmatic lipid droplets (CLDs; Murphy, 2012), although in some cases, such as in Chlamydomonas reinhardtii starchless mutants, they also accumulate in chloroplasts (Fan et al., 2011; Goodson et al., 2011). Recent studies indicate that the CLDs are closely associated with ER membranes and possibly, chloroplast envelope membranes as well (Goodson et al., 2011; Peled et al., 2012).Green microalgae also contain two distinct types of chloroplastic lipid droplets. The first type is plastoglobuli, similar in morphology to higher plants plastoglobuli (Bréhélin et al., 2007; Kessler and Vidi, 2007). The second type is the eyespot (stigma), part of the visual system in microalgae. The eyespot is composed of a cluster of β-carotene-containing lipid droplets organized in several layers between grana membranes in the chloroplast (Häder and Lebert, 2009; Kreimer, 2009). Recent proteomic analysis of algal eyespot proteins revealed that they contain diverse structural proteins, lipid and carotenoid metabolizing enzymes, transporters, and signal transduction components (Schmidt et al., 2006).The origin of TAG in microalgae is still not clear. In C. reinhardtii, it was found that the major fatty acids in the sn-2 position are 16:0, which according to the plant dogma, is made in the chloroplast (Fan et al., 2011). In C. reinhardtii, which lacks PC, monogalactosyldiacylglycerol (MGDG) was proposed to replace PC in the mobilization of fatty acids from plastidal galactoglycerolipids into TAG based on mutation of a galactoglycerolipid lipase (Li et al., 2012). Based on these results and others, it has been proposed that, in C. reinhardtii, triglycerides are primarily produced in the chloroplast or combined with ER (Li et al., 2012; Liu and Benning, 2013).Plants and algae lipid droplets contain structural major proteins localized at the lipid droplet periphery, and their major function seems to be stabilization and prevention of fusion (Huang, 1992, 1996; Katz et al., 1995; Frandsen et al., 2001; Liu et al., 2009). In plant seed oils, the major classes of lipid droplet proteins are oleosins and caleosins, which have a characteristic hydrophobic loop with a conserved three Pro domain (Hsieh and Huang, 2004; Capuano et al., 2007; Purkrtova et al., 2008; Tzen, 2012). Oleosin and caleosin analogs were also recently identified in some green microalgal species (Lin et al., 2012; Vieler et al., 2012; Huang et al., 2013). However, the most abundant lipid droplets proteins in green algae (Chloropyceae) are a new family of major lipid droplet proteins (MLDPs) structurally distinct from plant oleosins and caleosins (Moellering and Benning, 2010; Peled et al., 2011; Davidi et al., 2012). Plastoglobules have different major lipid-associated proteins termed plastoglobules-associated protein-fibrillins, which form a distinct protein family with no sequence or structural similarities to oleosins (Kim and Huang, 2003). We have previously identified in the plastoglobuli rich in β-carotene (βC-plastoglobuli) a lipid-associated protein termed carotene globule protein (CGP), whose degradation destabilized the lipid droplets (Katz et al., 1995). The proteome of C. reinhardtii lipid droplet indicates that algal CLDs also contain several enzymes, suggesting that they are involved in lipid metabolism (Nguyen et al., 2011).The halotolerant green algae Dunaliella bardawil and Dunaliella salina ‘Teodoresco’ are unique in that they accumulate under high light stress or nitrogen deprivation large amounts of plastidic lipid droplets (βC-plastoglobuli), which consist of TAG and two isomers of β-carotene, all trans and 9-cis (Ben-Amotz et al., 1982, 1988). D. bardawil also accumulates CLD under the same stress conditions, similar to other green algae (Davidi et al., 2012). It has been shown that the function of βC-plastoglobuli is to protect the photosynthetic system against photoinhibition (Ben-Amotz et al., 1989). The enzymatic pathway for β-carotene synthesis in D. bardawil and D. salina has been partly identified, but the subcellular localization of β-carotene biosynthesis is not known (Jin and Polle, 2009). The synthesis of β-carotene depends on TAG biosynthesis (Rabbani et al., 1998); however, the origin of βC-plastoglobuli is not known. Are they formed within the chloroplast, or are they made in the cytoplasm? Is the TAG in βC-plastoglobuli and CLD identical or different, and where is it formed?D. bardawil is an excellent model organism for isolation of lipid droplet for several reasons. First, D. bardawil contains large amounts of both CLD and βC-plastoglobuli (Ben-Amotz et al., 1982; Fried et al., 1982), making it possible to obtain sufficient amounts of proteins and lipids from the two types of lipid pools for detailed analyses. Second, Dunaliella do not have a rigid cell wall and can be lysed by a gentle osmotic shock, which does not rupture the chloroplast. Therefore, it is possible to sequentially release pure CLD and βC-plastoglobuli by a two-step lysis (Katz et al., 1995). Third, D. bardawil seems to lack the eyespot structure, which can be clearly observed in other Dunaliella spp. even in a light microscope or by electron microscopy, but has never been observed in D. bardawil by us. It avoids the risk of cross contamination of βC-plastoglobuli with eyespot proteins. Fourth, the availability of protein markers for the major lipid droplet-associated proteins, CGPs and MLDPs, enabled both good immunolocalization and careful monitoring of the purity of the preparations by western analysis.In this work, we describe the purification, lipid compositions, and protein profiles of two lipid pools from D. bardawil: CLD and plastidic βC-plastoglobuli. A detailed proteomic analysis of these lipid droplets will be described in another work. Combined with detailed electron microscopy studies, these results led to surprising conclusions regarding the origin of the plastidic βC-plastoglobuli.     

The Green Microalga Chlamydomonas reinhardtii Has a Single ω-3 Fatty Acid Desaturase That Localizes to the Chloroplast and Impacts Both Plastidic and Extraplastidic Membrane Lipids

The ω-3 polyunsaturated fatty acids account for more than 50% of total fatty acids in the green microalga Chlamydomonas reinhardtii, where they are present in both plastidic and extraplastidic membranes. In an effort to elucidate the lipid desaturation pathways in this model alga, a mutant with more than 65% reduction in total ω-3 fatty acids was isolated by screening an insertional mutant library using gas chromatography-based analysis of total fatty acids of cell pellets. Molecular genetics analyses revealed the insertion of a TOC1 transposon 113 bp upstream of the ATG start codon of a putative ω-3 desaturase (CrFAD7; locus Cre01.g038600). Nuclear genetic complementation of crfad7 using genomic DNA containing CrFAD7 restored the wild-type fatty acid profile. Under standard growth conditions, the mutant is indistinguishable from the wild type except for the fatty acid difference, but when exposed to short-term heat stress, its photosynthesis activity is more thermotolerant than the wild type. A comparative lipidomic analysis of the crfad7 mutant and the wild type revealed reductions in all ω-3 fatty acid-containing plastidic and extraplastidic glycerolipid molecular species. CrFAD7 was localized to the plastid by immunofluorescence in situ hybridization. Transformation of the crfad7 plastidial genome with a codon-optimized CrFAD7 restored the ω-3 fatty acid content of both plastidic and extraplastidic lipids. These results show that CrFAD7 is the only ω-3 fatty acid desaturase expressed in C. reinhardtii, and we discuss possible mechanisms of how a plastid-located desaturase may impact the ω-3 fatty acid content of extraplastidic lipids.Research on lipid metabolism in microalgae has flourished in recent years due to their potential as a rich source of ω-3 fatty acids (Guschina and Harwood, 2006; Khozin-Goldberg et al., 2011) and as a feedstock for biodiesel (Hu et al., 2008b; Rosenberg et al., 2008; Beer et al., 2009; Radakovits et al., 2010; Wijffels and Barbosa, 2010; Merchant et al., 2012; Work et al., 2012). Oils produced by microalgae resemble that of plants (Hu et al., 2008b), with the exception that they contain higher proportions of polyunsaturated fatty acid (PUFA) species (Harwood and Guschina, 2009). Desaturation of acyl groups in glycerolipids is catalyzed by fatty acid desaturases (FADs), which insert a C=C bond at a specifically defined position of an acyl chain (Shanklin and Cahoon, 1998). The degree of unsaturation of fatty acid components largely determines the chemical property and thus the utility of the oils produced. FADs have been one of the major tools for the genetic engineering of oil composition in land crops (Shanklin and Cahoon, 1998; Napier et al., 1999). In view of biodiesel applications, low PUFA content is advantageous in algal oil because of oxidation issues (Frankel, 1991).With the suites of sophisticated molecular genetic and genomic tools developed in the green microalga Chlamydomonas reinhardtii and the existence of substantial literature related to its cell biology, physiology, and biochemistry, this organism has emerged as a major model for research on algal oil (Radakovits et al., 2010; Merchant et al., 2012; Liu and Benning, 2013). Although the understanding of lipid metabolism in C. reinhardtii largely relies on sequence homologies to other models (Riekhof et al., 2005) and is still rather limited compared with the model plant Arabidopsis (Arabidopsis thaliana; Li-Beisson et al., 2010), functional studies based on mutants have started to provide important insights into the biosynthesis and turnover of membrane and storage lipids in this model alga (Riekhof et al., 2005; Work et al., 2010; Fan et al., 2011; Goodson et al., 2011; Boyle et al., 2012; Li et al., 2012a, 2012b; Yoon et al., 2012).In C. reinhardtii, C16 and C18 PUFAs (ω-3 + ω-6) make up to 60 mol% of total membrane fatty acids, of which more than 80% are ω-3 species (Giroud and Eichenberger, 1988; Siaut et al., 2011). Biochemical evidence for lipid-linked desaturation of fatty acyl chains has been established in C. reinhardtii over 20 years (Giroud and Eichenberger, 1989), but only two C. reinhardtii mutants affected in fatty acid desaturation have been described to date. These are crfad6 (hf-9), an insertional mutant for the plastidial ω-6 desaturase FAD6 (Sato et al., 1995), and microRNA-based silenced lines for the Δ4 desaturase CrΔ4FAD (Zäuner et al., 2012). The putative microsomal Δ12 desaturase FAD2 (Chi et al., 2008) and front-end ω-13 desaturase (Kajikawa et al., 2006) have been characterized by heterologous expression in the methylotrophic yeast Pichia pastoris, but no mutant is available. Moreover, although ω-3 PUFA is the most abundant fatty acid class in C. reinhardtii, the ω-3 desaturase remains uncharacterized, and no mutant with specific reduction in ω-3 content has been isolated so far.In Arabidopsis and C. reinhardtii, ω-3 PUFAs are present in both plastidic and extraplastidic lipids such as monogalactosyldiacylglycerol (MGDG) and phosphatidylethanolamine (PtdEtn), respectively (Mendiola-Morgenthaler et al., 1985; Giroud et al., 1988). While in plants there are distinct genes for plastidial and extraplastidial ω-3 FADs (Wallis and Browse, 2002), only one putative ω-3 desaturase seems encoded in the C. reinhardtii genome (version 5.0; Merchant et al., 2007). This raises several intriguing possibilities, including the existence of a mechanism to export ω-3 acyls from their site of biogenesis to other membranes or a dual localization of the ω-3 desaturase homolog (plastid and endoplasmic reticulum [ER]). In this study, we report the identification and characterization of a C. reinhardtii mutant defective in the promoter region of the putative ω-3 FAD encoded by the Cre01.g038600 locus. We show that while this enzyme is localized to plastids, impairment in its expression leads to a reduction of ω-3 fatty acids acylated to both plastidial and ER lipids. Additionally, using plastidial transformation of the mutant, it is demonstrated that the location of this desaturase in the plastid alone is sufficient to ensure normal ω-3 fatty acid content in extraplastidic lipids. Possible acyl desaturation and trafficking mechanisms implied by these findings are discussed.     

Putting the Squeeze on Plasmodesmata: A Role for Reticulons in Primary Plasmodesmata Formation

Primary plasmodesmata (PD) arise at cytokinesis when the new cell plate forms. During this process, fine strands of endoplasmic reticulum (ER) are laid down between enlarging Golgi-derived vesicles to form nascent PD, each pore containing a desmotubule, a membranous rod derived from the cortical ER. Little is known about the forces that model the ER during cell plate formation. Here, we show that members of the reticulon (RTNLB) family of ER-tubulating proteins in Arabidopsis (Arabidopsis thaliana) may play a role in the formation of the desmotubule. RTNLB3 and RTNLB6, two RTNLBs present in the PD proteome, are recruited to the cell plate at late telophase, when primary PD are formed, and remain associated with primary PD in the mature cell wall. Both RTNLBs showed significant colocalization at PD with the viral movement protein of Tobacco mosaic virus, while superresolution imaging (three-dimensional structured illumination microscopy) of primary PD revealed the central desmotubule to be labeled by RTNLB6. Fluorescence recovery after photobleaching studies showed that these RTNLBs are mobile at the edge of the developing cell plate, where new wall materials are being delivered, but significantly less mobile at its center, where PD are forming. A truncated RTNLB3, unable to constrict the ER, was not recruited to the cell plate at cytokinesis. We discuss the potential roles of RTNLBs in desmotubule formation.Plasmodesmata (PD), the small pores that connect higher plant cells, are complex structures of about 50 nm in diameter. Each PD pore is lined by the plasma membrane and contains an axial endoplasmic reticulum (ER)-derived structure known as the desmotubule (Overall and Blackman, 1996; Maule, 2008; Tilsner et al., 2011). The desmotubule is an enigmatic structure whose function has not been fully elucidated. The small spiraling space between the desmotubule and the plasma membrane, known as the cytoplasmic sleeve, is almost certainly a conduit for the movement of small molecules (Oparka et al., 1999). Some reports, however, suggest that the desmotubule may also function in cell-to-cell trafficking, providing an ER-derived pathway between cells along which macromolecules may diffuse (Cantrill et al., 1999). The desmotubule is one of the most tightly constricted membrane structures found in nature (Tilsner et al., 2011), but the forces that generate its intense curvature are not understood. In most PD, the desmotubule is a tightly furled tube of about 15 nm in diameter in which the membranes of the ER are in close contact along its length. The desmotubule may balloon out in the region of the middle lamella into a central cavity, but at the neck regions of the PD pore it is tightly constricted (Robinson-Beers and Evert, 1991; Ding et al., 1992; Glockmann and Kollmann, 1996; Overall and Blackman, 1996; Ehlers and Kollmann, 2001). Studies of PD using GFP targeted to the ER lumen (e.g. GFP-HDEL) have shown that GFP is excluded from the desmotubule due to the constriction of ER membranes in this structure (Oparka et al., 1999; Crawford and Zambryski, 2000; Martens et al., 2006; Guenoune-Gelbart et al., 2008). Therefore, lumenal GFP is unable to move between plant cells unless the membranes of the desmotubule become relaxed in some way. On the other hand, dyes and some proteins inserted into the ER membrane can apparently move through the desmotubule, either along the membrane or through the lumen, at least under some conditions (Grabski et al., 1993; Cantrill et al., 1999; Martens et al., 2006; Guenoune-Gelbart et al., 2008).Recently, a number of proteins have been described in mammalian, yeast, and plant systems that induce extreme membrane curvature. Among these are the RETICULONS (RTNs), integral membrane proteins that induce curvature of the ER to form tubules (Voeltz et al., 2006; Hu et al., 2008; Tolley et al., 2008, 2010; Sparkes et al., 2010). In animals, RTNs have been shown to be involved in a wide array of endomembrane-related processes, including intracellular transport and vesicle formation, and as RTNs can also influence axonal growth, they may have roles in neurodegenerative disorders such as Alzheimer’s disease (Yang and Strittmatter, 2007). Arabidopsis (Arabidopsis thaliana) has 21 RTN homologs, known as RTNLBs (Nziengui et al., 2007; Sparkes et al., 2010), considerably more than in yeast or mammals, but most have not been examined. RTNLBs contain two unusually long hydrophobic helices that form reentrant loops (Voeltz et al., 2006; Hu et al., 2008; Sparkes et al., 2010; Tolley et al., 2010). These are thought to induce membrane curvature by the molecular wedge principle (Hu et al., 2008; Shibata et al., 2009). When RTNLBs are overexpressed transiently in cells expressing GFP-HDEL, the ER becomes tightly constricted and GFP-HDEL is excluded from the lumen of the constricted ER tubules (Tolley et al., 2008, 2010), a situation similar to that which occurs in desmotubules (Oparka et al., 1999; Crawford and Zambryski, 2000; Martens et al., 2006). In vitro studies with isolated membranes have shown that the degree of tubulation is proportional to the number and spacing of RTNLB proteins in the membrane (Hu et al., 2008). For example, to constrict the ER membrane into a structure of 15 nm, the diameter of a desmotubule, would require RTNLBs to be inserted every 2 nm or less along the desmotubule axis (Hu et al., 2008), potentially making the desmotubule an extremely protein-rich structure (Tilney et al., 1991). Interestingly, a number of RTNLB proteins appear in the recently described PD proteome (Fernandez-Calvino et al., 2011), suggesting that RTNLBs are good candidates for proteins that model the cortical ER into desmotubules.Primary PD form at cytokinesis during the assembly of the cell plate (Hawes et al., 1981; Hepler, 1982). Of the numerous studies devoted to the structure of the cell plate, very few have examined the behavior of the ER during cytokinesis. During mitosis, elements of the ER are located in the spindle apparatus, separated from the cytoplasm (Hepler, 1980). Just prior to cytokinesis, there is a relative paucity of ER in the region destined to become the cell plate (Hepler, 1980; Hawes et al., 1981). The studies of Hawes et al. (1981) and Hepler (1982), exploiting heavy-metal impregnation of the ER, showed that during the formation of the new cell plate, strands of cortical ER are inserted across the developing wall, between the Golgi-derived vesicles that deposit wall materials. These ER strands become increasingly thinner during formation of the desmotubule, eventually excluding heavy metal stains from the ER lumen (Hepler, 1982). The center of the desmotubule often appears electron opaque in transmission electron microscopy images and has been referred to as the central rod (Overall and Blackman, 1996). This structure may consist of proteins that extend from the inner ER leaflets or may correspond to head groups of the membrane lipids themselves. In the fully formed primary PD, the desmotubule remains continuous with the cortical ER that runs close to the new cell wall (Hawes et al., 1981; Hepler, 1982; Oparka et al., 1994).Here, we show that two of the RTNLBs present in the PD proteome, RTNLB3 and RTNLB6, become localized to the cell plate during the formation of primary PD. These RTNLBs remain associated with the desmotubule in fully formed PD and are immobile, as evidenced by fluorescence recovery after photobleaching (FRAP) studies. A truncated version of RTNLB3, in which the second hydrophobic region was deleted (Sparkes et al., 2010), was not recruited to the cell plate at cytokinesis. We suggest that RTNLBs play an important role in the formation of primary PD and discuss mechanisms by which these proteins may model the ER into desmotubules.     

Control of Tobacco mosaic virus Movement Protein Fate by CELL-DIVISION-CYCLE Protein48

Like many other viruses, Tobacco mosaic virus replicates in association with the endoplasmic reticulum (ER) and exploits this membrane network for intercellular spread through plasmodesmata (PD), a process depending on virus-encoded movement protein (MP). The movement process involves interactions of MP with the ER and the cytoskeleton as well as its targeting to PD. Later in the infection cycle, the MP further accumulates and localizes to ER-associated inclusions, the viral factories, and along microtubules before it is finally degraded. Although these patterns of MP accumulation have been described in great detail, the underlying mechanisms that control MP fate and function during infection are not known. Here, we identify CELL-DIVISION-CYCLE protein48 (CDC48), a conserved chaperone controlling protein fate in yeast (Saccharomyces cerevisiae) and animal cells by extracting protein substrates from membranes or complexes, as a cellular factor regulating MP accumulation patterns in plant cells. We demonstrate that Arabidopsis (Arabidopsis thaliana) CDC48 is induced upon infection, interacts with MP in ER inclusions dependent on the MP N terminus, and promotes degradation of the protein. We further provide evidence that CDC48 extracts MP from ER inclusions to the cytosol, where it subsequently accumulates on and stabilizes microtubules. We show that virus movement is impaired upon overexpression of CDC48, suggesting that CDC48 further functions in controlling virus movement by removal of MP from the ER transport pathway and by promoting interference of MP with microtubule dynamics. CDC48 acts also in response to other proteins expressed in the ER, thus suggesting a general role of CDC48 in ER membrane maintenance upon ER stress.Plant viruses are obligate intracellular pathogens that replicate in association with host membranes (Laliberté and Sanfaçon, 2010) and subvert host intra- and intercellular trafficking pathways to achieve cell-to-cell and systemic spread (Harries and Ding, 2011; Niehl and Heinlein, 2011). In the case of the well-studied Tobacco mosaic virus (TMV), viral replication factories form on membranes of the endoplasmic reticulum (ER; Heinlein et al., 1995, 1998). As the plant ER is continuous between cells through plasmodesmata (PD; Ding et al., 1992), this membrane network provides a direct pathway for the spread of replicated virus from the replication sites in infected cells into the ER network of noninfected cells. The spread of plant viruses depends on virus-encoded movement proteins (MPs; Deom et al., 1987; Lucas, 2006). The MP of TMV facilitates the cell-to-cell passage of the infectious particle by forming a ribonucleoprotein complex with the viral RNA (Citovsky et al., 1990) and by increasing the size exclusion limit of PD (Wolf et al., 1989).During the course of infection, as well as when ectopically expressed, the MP associates with PD, the ER/actin network, and microtubules (Heinlein et al., 1995, 1998; Reichel and Beachy, 1998; Wright et al., 2007; Sambade et al., 2008; Hofmann et al., 2009; Boutant et al., 2010; Peña and Heinlein, 2012; Supplemental Fig. S1). Shortly after infection of a new cell, the MP localizes to small, mobile, ER-associated particles proposed to play a role in PD targeting of the viral RNA (Boyko et al., 2007; Sambade et al., 2008). Similar small, mobile MP particles are observed early upon ectopic expression of the protein. These particles colocalize with RNA and undergo stop-and-go movements in association with the ER (Sambade et al., 2008). The particle movements pause at microtubule proximal sites and their detachment requires microtubule polymerization (Sambade et al., 2008). These observations suggest that the interaction with the microtubule system plays a critical role in the maturation and ER-mediated delivery of infectious viral RNA particles to PD during early infection stages. Consistently, tobacco (Nicotiana tabacum) mutants with reduced microtubule dynamics exhibit reduced TMV movement (Ouko et al., 2010). Following virus movement, the previously infected cell further accumulates MP at the ER, a process that coincides with the formation of large ER inclusions that contain viral replicase and viral RNA in addition to MP and likely function as virus factories (Heinlein et al., 1998; Más and Beachy, 1999). In mature form, these inclusions may represent the so-called viroplasms or X-bodies described in the classical literature (Bawden and Sheffield, 1939; Esau and Cronshaw, 1967; Hills et al., 1987). Their formation is associated with rearrangements of the ER membrane and likely mediated by the accumulated MP since the inclusions diminish and reconstitute a native ER structure when MP becomes degraded by the 26S proteasome (Reichel and Beachy, 1998, 2000). Transfected cells accumulate MP in similar inclusions as those formed during infection, indicating that accumulated MP is indeed necessary and sufficient to form inclusions in association with the ER (Reichel and Beachy, 1998; Supplemental Fig. S1). Following accumulation of MP in virus factories, the infected cells accumulate the MP also along microtubules (Heinlein et al., 1998). The accumulation of MP in virus factories and on microtubules in cells behind the leading front of infection is dispensable for virus movement (Heinlein et al., 1998; Boyko et al., 2000a). At these late infection stages, the virus factories may enable the virus to produce high virion titers (Laliberté and Sanfaçon, 2010; Tilsner et al., 2012), and the subsequent accumulation along microtubules may play a role in withdrawing MP from the cell-to-cell communication pathway (Curin et al., 2007) and in stockpiling MP prior to degradation (Padgett et al., 1996; Gillespie et al., 2002).The molecular mechanisms that guide the MP to the ER and subsequently to microtubules during infection are not known. The MP is a hydrophobic protein that behaves like a membrane-integral or tightly membrane-associated protein in differential fractionation experiments and contains two predicted transmembrane domains (Reichel and Beachy, 1998; Brill et al., 2000, 2004) involved in ER association (Fujiki et al., 2006). The association with microtubules depends on MP amino acids 1 to 213 required for MP function (Kahn et al., 1998; Boyko et al., 2000b,Boyko et al., 2000c, 2002; Kotlizky et al., 2001). Moreover, certain amino acid exchange mutations known to affect the function of MP in virus movement in a temperature-sensitive manner also affect the ability of MP to interact with microtubules (Boyko et al., 2007,Boyko et al., 2000b). Interestingly, these mutations cluster together in a short domain of 25 amino acids showing a structural similarity with the M-loop of tubulin involved in tubulin-tubulin interactions (Boyko et al., 2000b; Waigmann et al., 2007). Importantly, this M-loop similarity domain overlaps with the predicted transmembrane domain (Brill et al., 2000, 2004) thus suggesting that the association of MP with membranes or microtubules is an alternative event that may depend on specific posttranslational modifications or specific folds of MP. However, although the different subcellular localizations of MPs during the course of infection indicate directional transport of MP from the ER to microtubules and may indicate different folds and functions of the protein when associated with these different subcellular components, the mechanism that controls the subcellular localization and, thus, the fate and function of MP is not known.Here, we identify CELL-DIVISION-CYCLE protein48 (CDC48), named p97/VCP (Valosin-containing protein) in mammals and Cdc48p in yeast (Saccharomyces cerevisiae), as a cellular factor regulating MP subcellular accumulation patterns. CDC48 functions are well characterized in mammalian and yeast systems but remain poorly investigated in plants. Yeast and mammalian CDC48s are essential, conserved chaperones involved in diverse cellular processes by controlling protein fate through extraction of substrates from membranes or complexes (Tsai et al., 2002; Meusser et al., 2005; Römisch, 2005; Rumpf and Jentsch, 2006; Schrader et al., 2009; Eisele et al., 2010; Meyer et al., 2012; Yamanaka et al., 2012). We show that virus infection leads to the induction of Arabidopsis (Arabidopsis thaliana) CDC48 isoforms and demonstrate a function of CDC48 in ER maintenance upon ER stress conditions. We further demonstrate that CDC48 interacts with MP and that CDC48 activity is required for MP degradation. Interaction of CDC48 with MP depends on the MP N terminus, which is required for degradation of the protein, for PD localization and microtubule accumulation of MP, and for function of MP in cell-to-cell transport of the viral RNA. Overexpressed CDC48 shifts MP subcellular localization from ER inclusions to microtubules, suggesting that CDC48 extracts the MP from ER-associated inclusions, where it accumulates in midstages of infection, to the cytosol, where it accumulates along microtubules during late infection stages. Moreover, overexpression of active, but not inactive, CDC48 inhibits virus movement. Our data demonstrate that a CDC48-dependent pathway leading to the clearance of ER-associated protein inclusions exists in plants, that plant viral MPs are substrates for this pathway, and that this pathway determines viral protein fate during infection. We suggest that CDC48-mediated extraction of MP from the ER is part of a plant defense response to remove MP from the ER, the compartment the virus uses for replication and movement.     

ROOT HAIR DEFECTIVE3 Family of Dynamin-Like GTPases Mediates Homotypic Endoplasmic Reticulum Fusion and Is Essential for Arabidopsis Development

In all eukaryotic cells, the endoplasmic reticulum (ER) forms a tubular network whose generation requires the fusion of ER membranes. In Arabidopsis (Arabidopsis thaliana), the membrane-bound GTPase ROOT HAIR DEFECTIVE3 (RHD3) is a potential candidate to mediate ER fusion. In addition, Arabidopsis has two tissue-specific isoforms of RHD3, namely RHD3-like (RL) proteins, and their function is not clear. Here, we show that a null allele of RHD3, rhd3-8, causes growth defects and shortened root hairs. A point mutant, rhd3-1, exhibits a more severe growth phenotype than the null mutant, likely because it exerts a dominant-negative effect on the RL proteins. Genetic analysis reveals that the double deletion of RHD3 and RL1 is lethal and that the rhd3 rl2 plants produce no viable pollen, suggesting that the RL proteins are redundant to RHD3. RHD3 family proteins can replace Sey1p, the homolog of RHD3 in yeast (Saccharomyces cerevisiae), in the maintenance of ER morphology, and they are able to fuse membranes both in vivo and in vitro. Our results suggest that RHD3 proteins mediate ER fusion and are essential for plant development and that the formation of the tubular ER network is of general physiological significance.In all eukaryotic cells, the endoplasmic reticulum (ER) comprises a continuous membrane system of sheets and tubules (Baumann and Walz, 2001; Shibata et al., 2006). ER tubules frequently connect through homotypic membrane fusion to form a reticular network (Lee and Chen, 1988; Prinz et al., 2000; Du et al., 2004). ER fusion in metazoans is mediated by the atlastins (ATLs), a class of dynamin-like, membrane-bound GTPases (Hu et al., 2009; Orso et al., 2009). ATL possesses a cytoplasmic N-terminal GTPase domain, followed by a helical domain, two closely spaced transmembrane domains, and a C-terminal cytosolic tail. ATL proteins localize mostly to ER tubules and they interact with the tubule-shaping proteins, reticulons and DP1 (Hu et al., 2009). A role for the ATLs in ER fusion is suggested by the fact that depletion of ATLs leads to long, nonbranched ER tubules in cultured cells (Hu et al., 2009) and to ER fragmentation in Drosophila melanogaster (Orso et al., 2009), possibly due to insufficient fusion between the tubules. Nonbranched ER tubules are also observed upon the expression of dominant-negative ATL mutants (Hu et al., 2009). In addition, antibodies to ATL inhibit ER network formation in Xenopus laevis egg extracts (Hu et al., 2009). Moreover, proteoliposomes containing purified D. melanogaster ATL undergo GTP-dependent fusion in vitro (Orso et al., 2009; Bian et al., 2011). The physiological significance of ER fusion is supported by the observation that mutations in human ATL1, the dominant isoform in the brain, cause hereditary spastic paraplegia (Zhao et al., 2001), a neurodegenerative disease characterized by axon shortening in corticospinal motor neurons and progressive spasticity and weakness of the lower limbs (Salinas et al., 2008).Many organisms lack ATL homologs. In yeast (Saccharomyces cerevisiae), another dynamin-like GTPase, Sey1p, has been found to share the same signature motifs and membrane topology as ATL (Hu et al., 2009). Recent work suggests that Sey1p mediates ER membrane fusion both in vivo and in vitro (Anwar et al., 2012). Cells lacking Sey1p grow normally (Hu et al., 2009), but additional mutation of an ER SNARE Ufe1p, which probably represents an alternative ER fusion mechanism in yeast, causes severe growth defects (Anwar et al., 2012). In Arabidopsis (Arabidopsis thaliana), the potential functional ortholog of ATL appears to be ROOT HAIR DEFECTIVE3 (RHD3; Hu et al., 2009), which was initially discovered by a genetic screen of root hair-defective mutants (Schiefelbein and Somerville, 1990). It is sequence related to Sey1p over the entire length (Wang et al., 1997; Brands and Ho, 2002). Mutations of RHD3 cause short and wavy root hairs (Schiefelbein and Somerville, 1990; Wang et al., 1997; Stefano et al., 2012) and defects in cell expansion (Wang et al., 2002).Despite the sequence homology between Sey1p and RHD3, it was reported that Sey1p could not replace RHD3 in plants and vice versa (Chen et al., 2011). Therefore, it is not clear whether RHD3 can mediate ER fusion. Another complication in plants is that the Arabidopsis RHD3 family also contains two RHD3-like (RL) proteins (Hu et al., 2003): RL1 is expressed only in pollen, whereas RL2 is expressed ubiquitously, but both are present at very low levels. Deletion of either RL protein causes no detectable defects in root hair development or overall growth (Chen et al., 2011). Whether RL proteins support the role of RHD3 in a tissue-specific manner remains to be investigated.Here, we have analyzed the function of RHD3 and RL proteins in Arabidopsis. We show that RHD3 and the two RL proteins play redundant roles but function during different stages of Arabidopsis development. In addition, we show that RHD3 proteins can functionally replace Sey1p in yeast and mediate ER membrane fusion.     

A New LxxxA Motif in the Transmembrane Helix3 of Maize Aquaporins Belonging to the Plasma Membrane Intrinsic Protein PIP2 Group Is Required for Their Trafficking to the Plasma Membrane

Aquaporins play important roles in maintaining plant water status under challenging environments. The regulation of aquaporin density in cell membranes is essential to control transcellular water flows. This work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) aquaporin subfamily, which is divided into two sequence-related groups (ZmPIP1s and ZmPIP2s). When expressed alone in mesophyll protoplasts, ZmPIP2s are efficiently targeted to the plasma membrane, whereas ZmPIP1s are retained in the endoplasmic reticulum (ER). A protein domain-swapping approach was utilized to demonstrate that the transmembrane domain3 (TM3), together with the previously identified N-terminal ER export diacidic motif, account for the differential localization of these proteins. In addition to protoplasts, leaf epidermal cells transiently transformed by biolistic particle delivery were used to confirm and refine these results. By generating artificial proteins consisting of a single transmembrane domain, we demonstrated that the TM3 of ZmPIP1;2 or ZmPIP2;5 discriminates between ER and plasma membrane localization, respectively. More specifically, a new LxxxA motif in the TM3 of ZmPIP2;5, which is highly conserved in plant PIP2s, was shown to regulate its anterograde routing along the secretory pathway, particularly its export from the ER.Aquaporins are of major importance to plant physiology, being essential for the regulation of transcellular water movement during growth and development (Maurel et al., 2008; Gomes et al., 2009; Heinen et al., 2009; Prado and Maurel, 2013; Chaumont and Tyerman, 2014). Aquaporins are small membrane proteins consisting of six transmembrane (TM) domains connected by five loops (A–E), and N and C termini facing the cytosol (Fig. 1A). They assemble as homotetramers and/or heterotetramers in the membrane, with each monomer acting as an independent water channel (Murata et al., 2000; Fetter et al., 2004; Gomes et al., 2009). Aquaporins form a highly divergent protein family in plants (Chaumont et al., 2001; Johanson et al., 2001), and this work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) family (Chaumont et al., 2001). The regulation of the subcellular localization of these proteins is a key process controlling their density in the plasma membrane (PM) and, hence, their physiological roles (Hachez et al., 2013).Open in a separate windowFigure 1.Swapping TM3 of ZmPIP2;5 with that of ZmPIP1;2 retains the protein in intracellular structures. A, Cartoons representing the chimeric proteins composed of ZmPIP2;5, in which each TM has been replaced by the corresponding TM from ZmPIP1;2. All proteins are drawn with the cytosolic domains facing down. ZmPIP2;5 and ZmPIP1;2 portions are shown in black and white, respectively. All chimeras were fused to the C terminus of mYFP, which is not displayed for clarity purposes. B, Confocal microscopy images of maize mesophyll protoplasts transiently coexpressing mYFP-tagged ZmPIP2;5-PIP1;2 TM chimeric proteins (green) and the ER marker mCFP:HDEL (cyan). FM4-64 was added as a PM marker (red). Arrowheads in image 13 indicate accumulation of the protein in punctate structures that are not labeled by mCFP:HDEL. The localization patterns of the proteins of interest are representative of a total of at least 22 cells from three independent experiments. C, Confocal microscopy images of a maize mesophyll protoplast transiently expressing mYFP:ZmPIP2;5-TM3_PIP1;2 (green) and ST:mCFP (magenta). Arrowheads indicate colocalization in Golgi stacks. The images are representative of a total of 17 cells from two independent experiments. Bar = 5 µm.PIP aquaporins cluster in two groups (PIP1s and PIP2s), which are highly conserved across species (Kammerloher et al., 1994; Chaumont et al., 2000, 2001; Johanson et al., 2001; Anderberg et al., 2012). We previously showed that the maize PIP1 and PIP2 isoforms exhibit different water channel activities when expressed in Xenopus laevis oocytes, with only PIP2s increasing the membrane water permeability coefficient (P_f; Chaumont et al., 2000). However, when ZmPIP1 and ZmPIP2 are coexpressed, the isoforms physically interact to modify their stability and trafficking to the oocyte membrane, and synergistically increase the oocyte P_f (Fetter et al., 2004). Similar synergistic interactions between PIP1s and PIP2s have been reported in numerous plant species (Temmei et al., 2005; Mahdieh et al., 2008; Vandeleur et al., 2009; Bellati et al., 2010; Ayadi et al., 2011; Horie et al., 2011; Yaneff et al., 2014).PIPs were originally thought to be exclusively localized in the PM and were named accordingly (Kammerloher et al., 1994). However, recent experiments have shown that not all PIPs are located to the PM under all conditions, and that regulation of PIP subcellular localization is a highly dynamic process involving protein interactions (Boursiac et al., 2005, 2008; Zelazny et al., 2007, 2009; Uehlein et al., 2008; Besserer et al., 2012; Luu et al., 2012). When expressed singly in maize leaf mesophyll protoplasts, fluorescently tagged ZmPIP1s and ZmPIP2s differ in their subcellular localization. ZmPIP1s are retained in the endoplasmic reticulum (ER), whereas ZmPIP2s are targeted to the PM (Zelazny et al., 2007). However, upon coexpression, ZmPIP1s are relocalized from the ER to the PM, where they perfectly colocalize with ZmPIP2s. This relocalization results from their physical interaction as demonstrated by Förster resonance energy transfer/fluorescence lifetime imaging microscopy and immunoprecipitation experiments (Zelazny et al., 2007). These results indicate that ZmPIP2s, but not ZmPIP1s, possess signals that allow them to be delivered to the PM, and that hetero-oligomerization is required for ZmPIP1 trafficking to the PM. Interestingly, a diacidic motif (DxE, Asp-any amino acid-Glu) located in the N terminus of ZmPIP2;4, ZmPIP2;5, and Arabidopsis (Arabidopsis thaliana) AtPIP2;1 was shown to be required to exit the ER (Zelazny et al., 2009; Sorieul et al., 2011). Diacidic motifs interact with Secretory protein24, which is thought to be the main cargo-selection protein of the Coat proteinII complex that mediates vesicle formation at ER export sites (Miller et al., 2003). However, not all PM-localized PIP2s contain a diacidic ER export signal (Zelazny et al., 2009). In addition, swapping the N-terminal region of ER-retained ZmPIP1;2 with that of PM-localized ZmPIP2;5, which contains the functional diacidic motif, is not sufficient to trigger ER export of the protein (Zelazny et al., 2009). This result suggests that other export signals might be present in PIP2s and/or ER retention signals might be present in PIP1s elsewhere than in the N terminus.To identify new signals regulating ZmPIP1 and ZmPIP2 protein trafficking along the secretory pathway, we used a protein domain swapping-based approach and identified the TM3 as an important region that discriminates between ER-retained ZmPIP1;2 and PM-localized ZmPIP2;5. Specific mutations in the TM3 region of ZmPIP2;5 allowed the identification of a new ZmPIP2-conserved LxxxA motif, which regulates its export from the ER.     

Arabidopsis sphingolipid fatty acid 2-hydroxylases (AtFAH1 and AtFAH2) are functionally differentiated in fatty acid 2-hydroxylation and stress responses

2-Hydroxy fatty acids (2-HFAs) are predominantly present in sphingolipids and have important physicochemical and physiological functions in eukaryotic cells. Recent studies from our group demonstrated that sphingolipid fatty acid 2-hydroxylase (FAH) is required for the function of Arabidopsis (Arabidopsis thaliana) Bax inhibitor-1 (AtBI-1), which is an endoplasmic reticulum membrane-localized cell death suppressor. However, little is known about the function of two Arabidopsis FAH homologs (AtFAH1 and AtFAH2), and it remains unclear whether 2-HFAs participate in cell death regulation. In this study, we found that both AtFAH1 and AtFAH2 had FAH activity, and the interaction with Arabidopsis cytochrome b₅ was needed for the sufficient activity. 2-HFA analysis of AtFAH1 knockdown lines and atfah2 mutant showed that AtFAH1 mainly 2-hydroxylated very-long-chain fatty acid (VLCFA), whereas AtFAH2 selectively 2-hydroxylated palmitic acid in Arabidopsis. In addition, 2-HFAs were related to resistance to oxidative stress, and AtFAH1 or 2-hydroxy VLCFA showed particularly strong responses to oxidative stress. Furthermore, AtFAH1 interacted with AtBI-1 via cytochrome b₅ more preferentially than AtFAH2. Our results suggest that AtFAH1 and AtFAH2 are functionally different FAHs, and that AtFAH1 or 2-hydroxy VLCFA is a key factor in AtBI-1-mediated cell death suppression.Sphingolipids are a large class of lipids present ubiquitously in eukaryotic cells and involved in various cellular processes such as signal transduction, protein transport, and programmed cell death as structural components of the plasma membrane and tonoplast and as signaling molecules (Dunn et al., 2004; Pata et al., 2010). These numerous functions are based on the structural diversity of sphingolipids. Complex sphingolipids are largely formed by a variety of head groups and a ceramide, which is comprised of a long-chain base amide linked to a fatty acid. One of the main characteristics of sphingolipid fatty acids is that large number of sphingolipids contained very-long-chain fatty acids (VLCFAs; fatty acids with ≥ C20; Raffaele et al., 2009). Another feature is the 2-hydroxylation of fatty acids (Alderson et al., 2005). The 2-hydroxy fatty acid (2-HFA)-containing sphingolipids are present in most organisms, including yeast (Saccharomyces cerevisiae), vertebrates, and plants. Especially in higher plants, 2-HFAs are contained in more than 90% complex sphingolipids such as glucosylceramides and glycosylinositolphosphorylceramides (Imai et al., 1995; Pata et al., 2010).The 2-hydroxylation of sphingolipid N-acyl chains is catalyzed in yeast and mammals by the enzyme sphingolipid fatty acid 2-hydroxylase (ScFAH1 and FA2H, respectively, and we refer to them as FAH in this report; Mitchell and Martin, 1997; Alderson et al., 2004; Hama, 2010). FAH is an endoplasmic reticulum (ER)-localized membrane protein composed of two domains: an N-terminal cytochrome b₅ (Cb5)-like domain and a putative catalytic site with the His motif (HXXHH), which coordinates the nonheme diiron cluster at the active site of the enzyme and is highly conserved among membrane-bound desaturases and hydroxylases. Mammalian FA2H is reported to catalyze the 2-hydroxylation of free fatty acid in vitro, which was dependent on a reconstituted electron transport system (Alderson et al., 2005). In mammals, FA2H-synthesized 2-HFAs abundantly accumulate in epidermal keratinocytes and the myelin-forming cells of the nervous system (Alderson et al., 2006; Uchida et al., 2007; Maldonado et al., 2008). Loss of FA2H in mouse altered the composition and physicochemical properties of sebum, causing alopecia (Maier et al., 2011). In addition, FA2H deficiency also causes a wide range of neurodegeneration phenotypes such as spastic paraplegia, leukodystrophy, and brain ion deposition (Schneider and Bhatia, 2010). Moreover, FA2H is a negative regulator of the cell cycle and facilitates dibutyryl-cyclic AMP-induced cell cycle exit in D6P2T schwannoma cells (Alderson and Hama, 2009). These studies suggest that 2-hydroxylation of fatty acids is important in various cellular functions in mammals. However, there had been few reports about plant FAHs in the past.Recently, we suggested that FAH could mediate the function of Bax inhibitor-1 (BI-1), which is an ER membrane-localized cell death suppressor widely conserved in animals and plants (Xu and Reed, 1998; Kawai-Yamada et al., 2001; Nagano et al., 2009). Arabidopsis (Arabidopsis thaliana) BI-1 (AtBI-1) plays important roles in the resistance to various biotic and abiotic stresses including pathogens, ER stress, ion stress, and oxidative stress generating reactive oxygen species (ROS; Kawai-Yamada et al., 2001, 2004, 2009; Watanabe and Lam, 2006, 2008; Ihara-Ohori et al., 2007; Ishikawa et al., 2010). Our recent work indicated that AtBI-1 interacted with ScFAH1, and that deletion of ScFAH1 prevented AtBI-1 from suppression of cell death induced by mammalian proapoptotic protein, Bax, in yeast cells (Nagano et al., 2009). In Arabidopsis, there are two highly identical FAH homologs, AtFAH1 and AtFAH2, which possess several His motifs like their yeast and mammalian counterparts. However, AtFAH1 and AtFAH2 lack a Cb5-like domain, which provides electrons to ScFAH1 and FA2H (Mitchell and Martin, 1997; Alderson et al., 2004; Nagano et al., 2009). Instead, we demonstrated the interaction between both AtFAHs and Arabidopsis Cb5s (AtCb5s), which are localized in the ER membrane or mitochondrial outer membrane, by using a split-ubiquitin yeast two-hybrid system. In addition, AtBI-1 also interacted with AtCb5s. Moreover, overexpression of AtBI-1 increased the amounts of 2-hydroxy VLCFAs. These results allowed us to speculate that AtBI-1 interacts with AtFAHs via AtCb5 to activate the synthesis of 2-hydroxy VLCFA in Arabidopsis, which leads to the suppression of cell death. However, it remains unclear whether 2-hydroxy VLCFA is related to the regulation of cell death or stress responses. Furthermore, little is known about the function of AtFAH1 and AtFAH2 in 2-HFA synthesis of plant cells.The results of this study confirmed that both AtFAHs are FAHs, and identified functional differences between AtFAH1 and AtFAH2 in fatty acid 2-hydroxylation in Arabidopsis. In addition, AtFAH1 modified the resistance to oxidative stress more strongly than AtFAH2, which was consistent with the preferential interaction with AtBI-1.