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.     

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.     

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.     

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.     

Composition,Assembly, and Trafficking of a Wheat Xylan Synthase Complex

Xylans play an important role in plant cell wall integrity and have many industrial applications. Characterization of xylan synthase (XS) complexes responsible for the synthesis of these polymers is currently lacking. We recently purified XS activity from etiolated wheat (Triticum aestivum) seedlings. To further characterize this purified activity, we analyzed its protein composition and assembly. Proteomic analysis identified six main proteins: two glycosyltransferases (GTs) TaGT43-4 and TaGT47-13; two putative mutases (TaGT75-3 and TaGT75-4) and two non-GTs; a germin-like protein (TaGLP); and a vernalization related protein (TaVER2). Coexpression of TaGT43-4, TaGT47-13, TaGT75-3, and TaGT75-4 in Pichia pastoris confirmed that these proteins form a complex. Confocal microscopy showed that all these proteins interact in the endoplasmic reticulum (ER) but the complexes accumulate in Golgi, and TaGT43-4 acts as a scaffold protein that holds the other proteins. Furthermore, ER export of the complexes is dependent of the interaction between TaGT43-4 and TaGT47-13. Immunogold electron microscopy data support the conclusion that complex assembly occurs at specific areas of the ER before export to the Golgi. A di-Arg motif and a long sequence motif within the transmembrane domains were found conserved at the NH₂-terminal ends of TaGT43-4 and homologous proteins from diverse taxa. These conserved motifs may control the forward trafficking of the complexes and their accumulation in the Golgi. Our findings indicate that xylan synthesis in grasses may involve a new regulatory mechanism linking complex assembly with forward trafficking and provide new insights that advance our understanding of xylan biosynthesis and regulation in plants.It is believed that Golgi-localized, multiprotein complexes synthesize plant hemicellulosic polysaccharides, including xylans. Such complexes are not well characterized in plants (Zeng et al., 2010; Atmodjo et al., 2011; Chou et al., 2012), which is in sharp contrast with mammalian and yeast cells (Jungmann and Munro, 1998; McCormick et al., 2000; Giraudo et al., 2001). Xylans are the most abundant plant hemicellulosic polysaccharides on Earth and play an important role in the integrity of cell walls, which is a key factor in plant growth. Any mutations affecting xylan backbone biosynthesis seem to result in abnormal growth of plants due mostly to thinning and weakening of secondary xylem walls, described as the irregular xylem (irx) phenotype. Thus, characterizing the xylan synthase complex (XSC) would have an impact on plant improvement, as well as many industrial applications related to food, feed, and biofuel production (Yang and Wyman, 2004; Faik, 2010). Although the Arabidopsis (Arabidopsis thaliana) irx mutants have revealed the involvement of several glycosyltransferase (GT) gene families in xylan biosynthesis (Brown et al., 2007, 2009; Lee et al., 2007, 2010; Wu et al., 2009, 2010), no XSCs have been purified/isolated from Arabidopsis tissues, and we still do not know whether some of the identified Arabidopsis GTs can assemble into functional XSCs. Furthermore, if GTs do assemble into XSCs, we don’t know the mechanisms by which plant cells control their assembly and cellular trafficking. In contrast to dicots, xylan synthase activity was recently immunopurified from etiolated wheat (Triticum aestivum) microsomes (Zeng et al., 2010). This purified wheat XS activity was shown to catalyze three activities, xylan-glucuronosyltransferase (XGlcAT), xylan-xylosyltransferase (XXylT), and xylan-arabinofuranosyltranferase (XAT), which work synergistically to synthesize xylan-type polymers in vitro (Zeng et al., 2008, 2010). This work focuses on describing protein composition, assembly, and trafficking of this purified wheat XS activity.In all eukaryotes, proteins of the secretory pathway (including GTs) are synthesized in the endoplasmic reticulum (ER) and modified as they go through the Golgi cisternae. Most proteins exit the ER from ER export sites (ERESs; Hanton et al., 2009) and use a signal-based sorting mechanism that allows them to be selectively recruited into vesicles coated by coat protein II complexes (Barlowe, 2003; Beck et al., 2008). For many Golgi-resident type II membrane proteins, di-Arg motifs, such as RR, RXR, and RRR located in their cytosolic NH₂-terminal ends, have been shown to be required for their ER export (Giraudo et al., 2003; Czlapinski and Bertozzi, 2006; Schoberer et al., 2009; Tu and Banfield, 2010). Interestingly, di-Arg motifs located ∼40 amino acids from the membrane on the cytosolic side can also be used to retrieve some type II ER-resident proteins from cis-Golgi (Schutze et al., 1994; Hardt et al., 2003; Boulaflous et al., 2009). In contrast to the signal-based sorting mechanism involved in trafficking between the ER and Golgi, the steady-state localization/retention of proteins (including GTs) in the Golgi is thought to occur through vesicular cycling. Cycling is influenced by various mechanisms, including the length and composition of the transmembrane domain (TMD) of type II GTs (Bretscher and Munro, 1993; Colley, 1997; van Vliet et al., 2003; Sousa et al., 2003; Sharpe et al., 2010), and the oligomerization/aggregation of GTs (kin hypothesis), which suggests that formation of homo- or heterooligomers of GTs in the Golgi may prevent their recruitment into clathrin-coated vesicles (Machamer, 1991; Nilsson et al., 1993; Weisz et al., 1993; Cole et al., 1996). Some Golgi-resident GTs are predicted to have a cleavable NH₂-terminal secretion signal peptide (SP) and would therefore exist as soluble proteins in the Golgi lumen. To maintain their proper Golgi localization, these processed GTs are likely part of multiprotein complexes anchored to integral membrane proteins. The fact that homologs of many of the trafficking proteins from mammalian and yeast cells are found in plants indicates that trafficking machineries of the plant secretory pathway are likely conserved (d’Enfert et al., 1992; Bar-Peled and Raikhel, 1997; Batoko et al., 2000; Pimpl et al., 2000; Phillipson et al., 2001; Hawes et al., 2008).It is becoming increasingly evident that understanding the mechanisms controlling protein-protein interaction, sorting, and trafficking of polysaccharide synthases (including XSCs) will help elucidate how plants regulate cell wall synthesis and deposition during their development. To this end, we believe that the purified wheat XS activity (Zeng et al., 2010) is an excellent model for this type of study. In this work, proteomics was used to determine the protein composition of the purified XS activity. Confocal microscopy and immunogold transmission electron microscopy (TEM) were used to investigate the assembly and trafficking of the complex. Our proteomics data showed that the purified activity contains two GTs, TaGT43-4 and TaGT47-13, two putative mutases, TaGT75-3 and TaGT75-4, and two non-GT proteins: a germin-like protein (TaGLP) belonging to cupin superfamily and a protein specific to monocots annotated as wheat vernalization-related protein 2 (TaVER2). Microscopy analyses revealed that all these proteins interact in the ER, but the assembled complexes accumulate in the Golgi. Export of these complexes from the ER is controlled by the interaction between TaGT43-4 and TaGT47-13. Characterization of the wheat XSC and its trafficking furthers our understanding of xylan biosynthesis in grasses and helps elucidate how polysaccharide synthase complexes are assembled, sorted, and maintained in different compartments of the secretory pathway.     

Spore Density Determines Infection Strategy by the Plant Pathogenic Fungus Plectosphaerella cucumerina

Necrotrophic and biotrophic pathogens are resisted by different plant defenses. While necrotrophic pathogens are sensitive to jasmonic acid (JA)-dependent resistance, biotrophic pathogens are resisted by salicylic acid (SA)- and reactive oxygen species (ROS)-dependent resistance. Although many pathogens switch from biotrophy to necrotrophy during infection, little is known about the signals triggering this transition. This study is based on the observation that the early colonization pattern and symptom development by the ascomycete pathogen Plectosphaerella cucumerina (P. cucumerina) vary between inoculation methods. Using the Arabidopsis (Arabidopsis thaliana) defense response as a proxy for infection strategy, we examined whether P. cucumerina alternates between hemibiotrophic and necrotrophic lifestyles, depending on initial spore density and distribution on the leaf surface. Untargeted metabolome analysis revealed profound differences in metabolic defense signatures upon different inoculation methods. Quantification of JA and SA, marker gene expression, and cell death confirmed that infection from high spore densities activates JA-dependent defenses with excessive cell death, while infection from low spore densities induces SA-dependent defenses with lower levels of cell death. Phenotyping of Arabidopsis mutants in JA, SA, and ROS signaling confirmed that P. cucumerina is differentially resisted by JA- and SA/ROS-dependent defenses, depending on initial spore density and distribution on the leaf. Furthermore, in situ staining for early callose deposition at the infection sites revealed that necrotrophy by P. cucumerina is associated with elevated host defense. We conclude that P. cucumerina adapts to early-acting plant defenses by switching from a hemibiotrophic to a necrotrophic infection program, thereby gaining an advantage of immunity-related cell death in the host.Plant pathogens are often classified as necrotrophic or biotrophic, depending on their infection strategy (Glazebrook, 2005; Nishimura and Dangl, 2010). Necrotrophic pathogens kill living host cells and use the decayed plant tissue as a substrate to colonize the plant, whereas biotrophic pathogens parasitize living plant cells by employing effector molecules that suppress the host immune system (Pel and Pieterse, 2013). Despite this binary classification, the majority of pathogenic microbes employ a hemibiotrophic infection strategy, which is characterized by an initial biotrophic phase followed by a necrotrophic infection strategy at later stages of infection (Perfect and Green, 2001). The pathogenic fungi Magnaporthe grisea, Sclerotinia sclerotiorum, and Mycosphaerella graminicola, the oomycete Phytophthora infestans, and the bacterial pathogen Pseudomonas syringae are examples of hemibiotrophic plant pathogens (Perfect and Green, 2001; Koeck et al., 2011; van Kan et al., 2014; Kabbage et al., 2015).Despite considerable progress in our understanding of plant resistance to necrotrophic and biotrophic pathogens (Glazebrook, 2005; Mengiste, 2012; Lai and Mengiste, 2013), recent debate highlights the dynamic and complex interplay between plant-pathogenic microbes and their hosts, which is raising concerns about the use of infection strategies as a static tool to classify plant pathogens. For instance, the fungal genus Botrytis is often labeled as an archetypal necrotroph, even though there is evidence that it can behave as an endophytic fungus with a biotrophic lifestyle (van Kan et al., 2014). The rice blast fungus Magnaporthe oryzae, which is often classified as a hemibiotrophic leaf pathogen (Perfect and Green, 2001; Koeck et al., 2011), can adopt a purely biotrophic lifestyle when infecting root tissues (Marcel et al., 2010). It remains unclear which signals are responsible for the switch from biotrophy to necrotrophy and whether these signals rely solely on the physiological state of the pathogen, or whether host-derived signals play a role as well (Kabbage et al., 2015).The plant hormones salicylic acid (SA) and jasmonic acid (JA) play a central role in the activation of plant defenses (Glazebrook, 2005; Pieterse et al., 2009, 2012). The first evidence that biotrophic and necrotrophic pathogens are resisted by different immune responses came from Thomma et al. (1998), who demonstrated that Arabidopsis (Arabidopsis thaliana) genotypes impaired in SA signaling show enhanced susceptibility to the biotrophic pathogen Hyaloperonospora arabidopsidis (formerly known as Peronospora parastitica), while JA-insensitive genotypes were more susceptible to the necrotrophic fungus Alternaria brassicicola. In subsequent years, the differential effectiveness of SA- and JA-dependent defense mechanisms has been confirmed in different plant-pathogen interactions, while additional plant hormones, such as ethylene, abscisic acid (ABA), auxins, and cytokinins, have emerged as regulators of SA- and JA-dependent defenses (Bari and Jones, 2009; Cao et al., 2011; Pieterse et al., 2012). Moreover, SA- and JA-dependent defense pathways have been shown to act antagonistically on each other, which allows plants to prioritize an appropriate defense response to attack by biotrophic pathogens, necrotrophic pathogens, or herbivores (Koornneef and Pieterse, 2008; Pieterse et al., 2009; Verhage et al., 2010).In addition to plant hormones, reactive oxygen species (ROS) play an important regulatory role in plant defenses (Torres et al., 2006; Lehmann et al., 2015). Within minutes after the perception of pathogen-associated molecular patterns, NADPH oxidases and apoplastic peroxidases generate early ROS bursts (Torres et al., 2002; Daudi et al., 2012; O’Brien et al., 2012), which activate downstream defense signaling cascades (Apel and Hirt, 2004; Torres et al., 2006; Miller et al., 2009; Mittler et al., 2011; Lehmann et al., 2015). ROS play an important regulatory role in the deposition of callose (Luna et al., 2011; Pastor et al., 2013) and can also stimulate SA-dependent defenses (Chaouch et al., 2010; Yun and Chen, 2011; Wang et al., 2014; Mammarella et al., 2015). However, the spread of SA-induced apoptosis during hyperstimulation of the plant immune system is contained by the ROS-generating NADPH oxidase RBOHD (Torres et al., 2005), presumably to allow for the sufficient generation of SA-dependent defense signals from living cells that are adjacent to apoptotic cells. Nitric oxide (NO) plays an additional role in the regulation of SA/ROS-dependent defense (Trapet et al., 2015). This gaseous molecule can stimulate ROS production and cell death in the absence of SA while preventing excessive ROS production at high cellular SA levels via S-nitrosylation of RBOHD (Yun et al., 2011). Recently, it was shown that pathogen-induced accumulation of NO and ROS promotes the production of azelaic acid, a lipid derivative that primes distal plants for SA-dependent defenses (Wang et al., 2014). Hence, NO, ROS, and SA are intertwined in a complex regulatory network to mount local and systemic resistance against biotrophic pathogens. Interestingly, pathogens with a necrotrophic lifestyle can benefit from ROS/SA-dependent defenses and associated cell death (Govrin and Levine, 2000). For instance, Kabbage et al. (2013) demonstrated that S. sclerotiorum utilizes oxalic acid to repress oxidative defense signaling during initial biotrophic colonization, but it stimulates apoptosis at later stages to advance necrotrophic colonization. Moreover, SA-induced repression of JA-dependent resistance not only benefits necrotrophic pathogens but also hemibiotrophic pathogens after having switched from biotrophy to necrotrophy (Glazebrook, 2005; Pieterse et al., 2009, 2012).Plectosphaerella cucumerina ((P. cucumerina, anamorph Plectosporum tabacinum) anamorph Plectosporum tabacinum) is a filamentous ascomycete fungus that can survive saprophytically in soil by decomposing plant material (Palm et al., 1995). The fungus can cause sudden death and blight disease in a variety of crops (Chen et al., 1999; Harrington et al., 2000). Because P. cucumerina can infect Arabidopsis leaves, the P. cucumerina-Arabidopsis interaction has emerged as a popular model system in which to study plant defense reactions to necrotrophic fungi (Berrocal-Lobo et al., 2002; Ton and Mauch-Mani, 2004; Carlucci et al., 2012; Ramos et al., 2013). Various studies have shown that Arabidopsis deploys a wide range of inducible defense strategies against P. cucumerina, including JA-, SA-, ABA-, and auxin-dependent defenses, glucosinolates (Tierens et al., 2001; Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014), callose deposition (García-Andrade et al., 2011; Gamir et al., 2012, 2014; Sánchez-Vallet et al., 2012), and ROS (Tierens et al., 2002; Sánchez-Vallet et al., 2010; Barna et al., 2012; Gamir et al., 2012, 2014; Pastor et al., 2014). Recent metabolomics studies have revealed large-scale metabolic changes in P. cucumerina-infected Arabidopsis, presumably to mobilize chemical defenses (Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014). Furthermore, various chemical agents have been reported to induce resistance against P. cucumerina. These chemicals include β-amino-butyric acid, which primes callose deposition and SA-dependent defenses, benzothiadiazole (BTH or Bion; Görlach et al., 1996; Ton and Mauch-Mani, 2004), which activates SA-related defenses (Lawton et al., 1996; Ton and Mauch-Mani, 2004; Gamir et al., 2014; Luna et al., 2014), JA (Ton and Mauch-Mani, 2004), and ABA, which primes ROS and callose deposition (Ton and Mauch-Mani, 2004; Pastor et al., 2013). However, among all these studies, there is increasing controversy about the exact signaling pathways and defense responses contributing to plant resistance against P. cucumerina. While it is clear that JA and ethylene contribute to basal resistance against the fungus, the exact roles of SA, ABA, and ROS in P. cucumerina resistance vary between studies (Thomma et al., 1998; Ton and Mauch-Mani, 2004; Sánchez-Vallet et al., 2012; Gamir et al., 2014).This study is based on the observation that the disease phenotype during P. cucumerina infection differs according to the inoculation method used. We provide evidence that the fungus follows a hemibiotrophic infection strategy when infecting from relatively low spore densities on the leaf surface. By contrast, when challenged by localized host defense to relatively high spore densities, the fungus switches to a necrotrophic infection program. Our study has uncovered a novel strategy by which plant-pathogenic fungi can take advantage of the early immune response in the host plant.     

The Endoplasmic Reticulum Is a Reservoir for WAVE/SCAR Regulatory Complex Signaling in the Arabidopsis Leaf

During plant cell morphogenesis, signal transduction and cytoskeletal dynamics interact to locally organize the cytoplasm and define the geometry of cell expansion. The WAVE/SCAR (for WASP family verprolin homologous/suppressor of cyclic AMP receptor) regulatory complex (W/SRC) is an evolutionarily conserved heteromeric protein complex. Within the plant kingdom W/SRC is a broadly used effector that converts Rho-of-Plants (ROP)/Rac small GTPase signals into Actin-Related Protein2/3 and actin-dependent growth responses. Although the components and biochemistry of the W/SRC pathway are well understood, a basic understanding of how cells partition W/SRC into active and inactive pools is lacking. In this paper, we report that the endoplasmic reticulum (ER) is an important organelle for W/SRC regulation. We determined that a large intracellular pool of the core W/SRC subunit NAP1, like the known positive regulator of W/SRC, the DOCK family guanine nucleotide-exchange factor SPIKE1 (SPK1), localizes to the surface of the ER. The ER-associated NAP1 is inactive because it displays little colocalization with the actin network, and ER localization requires neither activating signals from SPK1 nor a physical association with its W/SRC-binding partner, SRA1. Our results indicate that in Arabidopsis (Arabidopsis thaliana) leaf pavement cells and trichomes, the ER is a reservoir for W/SRC signaling and may have a key role in the early steps of W/SRC assembly and/or activation.The W/SRC (for WASP family verprolin homologous/suppressor of cAMP receptor regulatory complex) and Actin-Related Protein (ARP)2/3 complex are part of an evolutionarily conserved Rho-of-Plants (ROP)/Rac small GTPase signal transduction cascade that controls actin-dependent morphogenesis in a wide variety of tissues and developmental contexts (Smith and Oppenheimer, 2005; Szymanski, 2005; Yalovsky et al., 2008). Many of the components and regulatory relationships among the complexes were discovered based on the stage-specific cell-swelling and -twisting phenotypes of the distorted class of Arabidopsis (Arabidopsis thaliana) trichome mutants (Szymanski et al., 1999; Zhang et al., 2005, 2008; Djakovic et al., 2006; Le et al., 2006; Uhrig et al., 2007). However, in both maize (Zea mays) and Arabidopsis, W/SRC and/or ARP2/3 are required for normal pavement cell morphogenesis (Frank and Smith, 2002; Mathur et al., 2003b; Brembu et al., 2004). Compared with other Arabidopsis pavement cell mutants, the shape defects of the distorted group are relatively mild. However, the distorted mutants and spike1 (spk1) differ from most other morphology mutants in that they display gaps in the shoot epidermis, most frequently at the interface of pavement cells and stomata (Qiu et al., 2002; Le et al., 2003; Li et al., 2003; Mathur et al., 2003b; Zhang et al., 2005; Djakovic et al., 2006). The cell gaps may reflect either uncoordinated growth between neighboring cells or defective cortical actin-dependent secretion of polysaccharides and/or proteins that promote cell-cell adhesion (Smith and Oppenheimer, 2005; Hussey et al., 2006; Leucci et al., 2007).In tip-growing cells, there is a strict requirement for actin to organize the trafficking and secretion activities of the cell to restrict growth to the apex. In Arabidopsis, the W/SRC-ARP2/3 pathway is not an essential tip growth component, because null alleles of both W/SRC and ARP2/3 subunits do not cause noticeable pollen tube or root hair phenotypes (Le et al., 2003; Djakovic et al., 2006). However, reverse genetic analysis of the W/SRC subunit BRK1 and ARP2/3 in the tip-growing protonemal cells of Physcomitrella patens revealed the obvious importance of this pathway (Harries et al., 2005; Perroud and Quatrano, 2008). Along similar lines, in two different legume species, W/SRC subunits are required for a normal root nodulation response to symbiotic bacteria (Yokota et al., 2009; Miyahara et al., 2010), indicating a conditional importance for this pathway in root hair growth. These genetic studies centered on the W/SRC and ARP2/3 pathways, in addition to those that involve a broader collection of actin-based morphology mutants (Smith and Oppenheimer, 2005; Blanchoin et al., 2010), are defining important cytoskeletal proteins and new interactions with the endomembrane system during morphogenesis. However, it is not completely clear how unstable actin filaments and actin bundle networks dictate the growth patterns of cells (Staiger et al., 2009).The difficulty of understanding the functions of specific actin arrays can be explained, in part, by the fact that plant cells that employ a diffuse growth mechanism have highly unstable cortical actin filaments and large actin bundles that do not have a geometry that obviously relates to the direction of growth or a specific subcellular activity (Blanchoin et al., 2010). This is in contrast to the cortical endocytic actin patches in yeast (Saccharomyces cerevisiae; Evangelista et al., 2002; Kaksonen et al., 2003) and cortical meshworks in the lamellipodia of crawling cells (Pollard and Borisy, 2003) that reveal subcellular locations where actin works to locally control membrane dynamics. In thick-walled plant cells, the magnitude of the forces that accompany turgor-driven cell expansion exceed those that could be generated by actin polymerization by orders of magnitude (Szymanski and Cosgrove, 2009). Localized cell wall loosening or the assembly of an anisotropic cell wall generates asymmetric yielding responses to turgor-induced stress (Baskin, 2005; Cosgrove, 2005). Therefore, the actin-based control of cell boundary dynamics is indirect, and the actin cytoskeleton influences cell shape change, in part, by actin and/or myosin-dependent trafficking of hormone transporters (Geldner et al., 2001) and organelles (Prokhnevsky et al., 2008), including those that control the localized delivery of protein complexes and polysaccharides that pattern the cell wall (Leucci et al., 2007; Gutierrez et al., 2009). In this scheme for actin-based growth control, the actin network dynamically rearranges at spatial scales that span from approximately 1- to 10-µm subcellular domains that may locally position organelles (Cleary, 1995; Gibbon et al., 1999; Szymanski et al., 1999) to the more than 100-µm actin bundle networks that operate at the spatial scales of entire cells (Gutierrez et al., 2009; Dyachok et al., 2011). It is clear from the work of several laboratories that the W/SRC and ARP2/3 protein complexes are required to organize cortical actin and actin bundle networks in trichomes (Szymanski et al., 1999; Le et al., 2003; Deeks et al., 2004; Zhang et al., 2005) and cylindrical epidermal cells (Mathur et al., 2003b; Dyachok et al., 2008, 2011). A key challenge now is to understand how plant cells deploy these approximately 10- to 20-nm heteromeric protein complexes to influence the patterns of growth at cellular scales.The genetic and biochemical control of ARP2/3 is complicated, but this is a tractable problem in plants, because the pathway is relatively simple compared with most other species in which it has been characterized. For example, in organisms ranging from yeast to humans, there are multiple types of ARP2/3 activators, protein complexes, and pathways that activate ARP2/3 (Welch and Mullins, 2002; Derivery and Gautreau, 2010). However, the maize and Arabidopsis genomes encode only WAVE/SCAR homologous proteins that can potently activate ARP2/3 (Frank et al., 2004; Basu et al., 2005). Detailed genetic and biochemical analyses of the WAVE/SCAR gene family in Arabidopsis demonstrated that the plant activators function interchangeably within the context of the W/SRC and define the lone pathway for ARP2/3 activation (Zhang et al., 2008). Bioinformatic analyses are consistent with a prominent role for W/SRC in the angiosperms, because in general, WASH complex subunits, which are structurally similar to WAVE/SCAR proteins, are largely absent from the higher plant genomes, while WAVE/SCAR genes are highly conserved (Kollmar et al., 2012).The components and regulatory schemes of the W/SRC-ARP2/3 pathway in Arabidopsis and P. patens are conserved compared with vertebrate species that employ these same protein complexes (Szymanski, 2005). For example, mutant complementation tests indicate that human W/SRC and ARP2/3 complex subunits can substitute for the Arabidopsis proteins (Mathur et al., 2003b). Furthermore, biochemical assays of Arabidopsis W/SRC (Basu et al., 2004; El-Assal et al., 2004; Frank et al., 2004; Le et al., 2006; Uhrig et al., 2007) and ARP2/3 assembly (Kotchoni et al., 2009) have shown that the binary interactions among W/SRC subunits and ARP2/3 complex assembly mechanisms are indistinguishable from those that have been observed for human W/SRC (Gautreau et al., 2004) and yeast ARP2/3 (Winter et al., 1999). After an initial period of controversy concerning the biochemical control of W/SRC, it is now apparent that vertebrate W/SRC (Derivery et al., 2009; Ismail et al., 2009), like the ARP2/3 complex (Machesky et al., 1999), is intrinsically inactive and requires positive regulation by Rac and other factors to fully activate ARP2/3 (Ismail et al., 2009; Lebensohn and Kirschner, 2009; Chen et al., 2010). Although overexpression of dominant negative ROP mutants causes trichome swelling and a reduced trichome branch number (Fu et al., 2002), the involvement of ROPs in trichome morphogenesis has been difficult to prove with a loss-of-function ROP allele because so many ROPs are expressed in this cell type (Marks et al., 2009). Existing reports on ROP loss-of-function mutants demonstrate the importance of pavement cell morphogenesis but do not document a trichome phenotype (Fu et al., 2005; Xu et al., 2010). A recent report describes a clever strategy to generate ROP loss-of-function lines that used the ectopic expression of ROP-specific bacterial toxins. There was a strong association between inducible expression of the toxins and the appearance of trichomes with severe trichome swelling and reduced branch number phenotypes (Singh et al., 2012). Although the exact mechanism of ROP-dependent control of W/SRC remains to be determined, the results described above in combination with the detection of direct interactions between the ROPGEF SPK1, active forms of ROP, and W/SRC subunits (Basu et al., 2004, 2008; Uhrig et al., 2007) strongly suggest that W/SRC is a ROP effector complex.The major challenge in the field now is to better understand the cellular control of W/SRC and how the complex is partitioned into active and inactive pools. In mammalian cells that crawl on a solid substrate, current models propose that a cytosolic pool of inactive WAVE/SCAR proteins and W/SRC is locally recruited and activated at specific plasma membrane surfaces in response to signals from some unknown Rac guanine nucleotide-exchange factor (GEF), protein kinase, and/or lipid kinase (Oikawa et al., 2004; Lebensohn and Kirschner, 2009; Chen et al., 2010). However, in Drosophila melanogaster neurons (Bogdan and Klämbt, 2003) and cultured human melanoma cells (Steffen et al., 2004), there are large pools of W/SRC with a perinuclear or organelle-like punctate localization that has no obvious relationship to cell shape or motility, raising uncertainty about the cellular mechanisms of W/SRC activation and the importance of different subcellular pools of the complex.In plants, cell fractionation experiments indicate that SCAR1 and ARP2/3 have an increased association with membranes compared with their animal counterparts (Dyachok et al., 2008; Kotchoni et al., 2009). In tip-growing moss protonemal cells, both the W/SRC subunit BRK1 and ARP2/3 localize to a population of unidentified organelles within the apical zone (Perroud and Quatrano, 2008). Similar live-cell imaging experiments in Arabidopsis reported a plasma membrane localization for SCAR1 and BRK1 in a variety of shoot epidermal and root cortex, and their accumulation at young trichome branch tips and at three-way cell wall junctions may define subcellular domains for W/SRC-ARP2/3-dependent actin filament nucleation at the plasma membrane (Dyachok et al., 2008). However, to our knowledge, active W/SRC, defined here as the fraction of W/SRC that colocalizes with ARP2/3 or actin, has not been reported in plants, and the plasma membrane is not necessarily the only organelle involved in W/SRC regulation. For example, the reported accumulation of BRK1 and SCAR1 at three-way cell wall junctions has a punctate appearance at the cell cortex that may not simply correspond to the plasma membrane (Dyachok et al., 2008). Also, in young stage 4 trichomes, there was an uncharacterized pool of intracellular SCAR1, but not BRK1, that localized to relatively large punctate structures (Dyachok et al., 2008). The endoplasmic reticulum (ER) may also be involved in W/SRC regulation. The ER-localized DOCK family ROPGEF SPK1 (Zhang et al., 2010) physically associates with multiple W/SRC proteins (Uhrig et al., 2007; Basu et al., 2008) and, based on genetic criteria, is an upstream, positive regulator of the W/SRC-ARP2/3 pathway (Basu et al., 2008). In the leaf, one function of SPK1 is to promote normal trafficking between the ER and Golgi; however, arp2/3 mutants do not share ER-stress phenotypes with spk1 (Zhang et al., 2010), making it unclear if SPK1 and the ER are directly involved in W/SRC signaling.This paper focuses on the localization and control of the W/SRC subunit NAP1/GNARLED/NAPP/HEM1/2. Arabidopsis NAP1 directly interacts with the ROP/Rac effector subunit SRA1/PIROGI/KLUNKER/PIRP (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). Based on the equally severe syndrome of nap1 and arp2/3 null phenotypes, and double mutant analyses, the only known function of NAP1 is to positively regulate ARP2/3 (Brembu et al., 2004; Deeks et al., 2004; El-Din El-Assal et al., 2004; Li et al., 2004). The vertebrate SRA1-NAP1 dimer is important for W/SRC assembly (Gautreau et al., 2004) and forms an extended physical surface that trans-inhibits the C-terminal ARP2/3-activating domain of WAVE/SCAR (Chen et al., 2010). The plant NAP1 and SRA1 proteins share end-to-end amino acid conservation with their vertebrate homologs and may form a heterodimer with similar functions (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). We report here that Arabidopsis NAP1 is strongly associated with ER membranes. In a detailed series of localization experiments, we detect a complicated intracellular distribution of NAP1 among the ER, the nucleus, and unidentified submicrometer punctae. A large pool of ER-associated NAP1 is inactive, based on the low level of colocalization with actin.Its accumulation on the ER does not require activating signals from either SPK1 or SRA1. These data indicate that the ER is a reservoir for W/SRC signaling and suggest that early steps in the positive regulation of NAP1 and the W/SRC occur on the ER surface.     

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.     

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.