The Cellulase KORRIGAN Is Part of the Cellulose Synthase Complex

Plant growth and organ formation depend on the oriented deposition of load-bearing cellulose microfibrils in the cell wall. Cellulose is synthesized by a large relative molecular weight cellulose synthase complex (CSC), which comprises at least three distinct cellulose synthases. Cellulose synthesis in plants or bacteria also requires the activity of an endo-1,4-β-d-glucanase, the exact function of which in the synthesis process is not known. Here, we show, to our knowledge for the first time, that a leaky mutation in the Arabidopsis (Arabidopsis thaliana) membrane-bound endo-1,4-β-d-glucanase KORRIGAN1 (KOR1) not only caused reduced CSC movement in the plasma membrane but also a reduced cellulose synthesis inhibitor-induced accumulation of CSCs in intracellular compartments. This suggests a role for KOR1 both in the synthesis of cellulose microfibrils and in the intracellular trafficking of CSCs. Next, we used a multidisciplinary approach, including live cell imaging, gel filtration chromatography analysis, split ubiquitin assays in yeast (Saccharomyces cerevisiae NMY51), and bimolecular fluorescence complementation, to show that, in contrast to previous observations, KOR1 is an integral part of the primary cell wall CSC in the plasma membrane.Cellulose microfibrils are synthesized by a hexameric multiprotein complex at the plasma membrane called the cellulose synthase complex (CSC). Genetic analysis, expression data, and coimmunoprecipitation experiments have demonstrated that a functional CSC contains at least three different nonredundant cellulose synthase (CESA) isoforms (Höfte et al., 2007). CESA1, CESA3, and CESA6-like are involved in cellulose biosynthesis during primary cell wall deposition, whereas CESA4, CESA7, and CESA8 are essential for cellulose synthesis in the secondary cell wall (Taylor et al., 1999, 2000, 2003; Desprez et al., 2007; Persson et al., 2007). CSCs labeled by fluorescently tagged CESA proteins migrate in the plasma membrane along cortical microtubules (CMTs), propelled by the polymerization of the β-1,4-glucans (Paredez et al., 2006). Partial depolymerization of CMTs using oryzalin showed that the organized trajectories of CSCs depend on the presence of an intact CMT array. The CSC-microtubule interaction is mediated at least in part by a large protein, POMPOM2/CELLULOSE SYNTHASE INTERACTING1, that binds to both CESAs and microtubules (Lei et al., 2014). Interestingly, complete depolymerization of CMTs does not alter the velocity of the complexes, illustrating that CMTs are necessary for the guidance of CSCs but not for their movement (Paredez et al., 2006). The microtubule cytoskeleton also has a role in the secretion and internalization of CSCs (Crowell et al., 2009; Gutierrez et al., 2009)KORRIGAN1 (KOR1) is a membrane-bound endo-1,4-β-d-glucanase (EGase) that is also required for cellulose synthesis (Nicol et al., 1998). Enzymatic analysis of a recombinant and soluble form of the Brassica napus KOR1 homolog showed substrate specificity for low-substituted carboxymethyl cellulose and amorphous cellulose but no activity on crystalline cellulose, xyloglucans, or short cellulose oligomers (Mølhøj et al., 2001; Master et al., 2004). Fractionation of microsomes demonstrated that KOR1 is primarily present in plasma membrane fractions but also at low levels in a tonoplast-enriched fraction (Nicol et al., 1998). Similarly, the KOR1 ortholog from tomato (Solanum lycopersicum) was found in the plasma membrane and fractions enriched for the Golgi apparatus (Brummell et al., 1997). A GFP-KOR1 fusion protein expressed with the Cauliflower mosaic virus 35S promoter accumulated in the Golgi apparatus and post-Golgi compartments and the tonoplast (Robert et al., 2005). Surprisingly for an enzyme involved in cellulose synthesis, the protein could not be detected at the plasma membrane. Using this construct, it was also shown that KOR1 undergoes regulated intracellular cycling (Robert et al., 2005).Although numerous genetic studies indicate that KOR1 is required for cellulose synthesis in primary and secondary cell walls and during cell plate formation (Nicol et al., 1998; Peng et al., 2000; Zuo et al., 2000; Lane et al., 2001; Sato et al., 2001; Szyjanowicz et al., 2004), its precise role in the cellulose synthesis process remains unclear. It has been suggested that KOR1 might be a component of the CSC (Read and Bacic, 2002). However, until now there has been no experimental evidence for this in Arabidopsis (Arabidopsis thaliana), either with coprecipitation experiments or with localization studies (Szyjanowicz et al., 2004; Robert et al., 2005; Desprez et al., 2007). Numerous hypotheses have been proposed to explain the paradoxical role of KOR1 in cellulose synthesis (Robert et al., 2004). KOR1 might have a proofreading activity involved in hydrolyzing disordered amorphous cellulose to relieve stress generated during the assembly of glucan chains in cellulose microfibrils (Mølhøj et al., 2002). Alternatively, KOR1 may determine the length of individual cellulose chains, either during cellulose synthesis or once the microfibril has been incorporated in the wall. A third hypothesis is that KOR1 releases the cellulose microfibril from the CSC before the complex is internalized from the plasma membrane (Somerville, 2006). Studies in cotton (Gossypium hirsutum) fiber extracts identified sitosterol glucoside as a primer for the cellulose synthesis and suggested that KOR1 could be involved in their cleavage from the nascent glucan chain (Peng et al., 2002). However, this scenario is unlikely, since, at least for the bacterial CESA, which is homologous to plant CESAs, there is no evidence for the existence of lipid-linked precursors, as shown by the three-dimensional structure of an active complex (Morgan et al., 2013).In this study, we first confirmed previous observations (Paredez et al., 2008) that, in the leaky kor1-1 mutant, the velocity of the CSCs is reduced compared with that in a wild-type background but that, in addition, the mutation affects the ability of the cellulose synthesis inhibitor CGA325′615 (hereafter referred to as CGA) to induce the accumulation of GFP-CESA3 in a microtubule-associated compartment (MASC/small compartments carrying cellulose synthase complexes [SmaCCs]; Crowell et al., 2009; Gutierrez et al., 2009). This indicates that KOR1 plays a role both in the synthesis of cellulose and in the intracellular trafficking of the CSC. Using gel filtration approaches, we identified KOR1 in fractions of high molecular mass, suggesting that KOR1 is present in membranes as part of a protein complex. We next analyzed the dynamics of GFP-KOR1 expressed in the kor1-1 mutant background under the control of its endogenous promoter. GFP-KOR1 is found in discrete particles at the plasma membrane in the same cells as GFP-CESAs (Crowell et al., 2009). GFP-KOR1 plasma membrane particles migrate along linear trajectories with comparable velocities to those observed for GFP-CESAs. The organization of GFP-KOR1 at the plasma membrane also requires the presence of an intact microtubule array, suggesting that KOR1 and CESA trajectories in the plasma membrane are regulated in the same manner. GFP-KOR1 and mCherry-CESA1 partially colocalize in the plasma membrane, Golgi, and post-Golgi compartments. Finally, we provide evidence for direct interaction between KOR1 and primary cell wall CESA proteins using the membrane-based yeast (Saccharomyces cerevisiae NMY51) two-hybrid (MbYTH) system (Timmers et al., 2009) and bimolecular fluorescence complementation (BiFC). Our data support a new model in which KOR1 is an integral part of the CSC, where it plays a role not only in the synthesis of cellulose but also in the intracellular trafficking of the CSC.     

The Endocytosis of Cellulose Synthase in Arabidopsis Is Dependent on μ2, a Clathrin-Mediated Endocytosis Adaptin

Clathrin-mediated endocytosis (CME) is the best-characterized type of endocytosis in eukaryotic cells. Plants appear to possess all of the molecular components necessary to carry out CME; however, functional characterization of the components is still in its infancy. A yeast two-hybrid screen identified μ2 as a putative interaction partner of CELLULOSE SYNTHASE6 (CESA6). Arabidopsis (Arabidopsis thaliana) μ2 is homologous to the medium subunit 2 of the mammalian ADAPTOR PROTEIN COMPLEX2 (AP2). In mammals, the AP2 complex acts as the central hub of CME by docking to the plasma membrane while concomitantly recruiting cargo proteins, clathrin triskelia, and accessory proteins to the sites of endocytosis. We confirmed that μ2 interacts with multiple CESA proteins through the μ-homology domain of μ2, which is involved in specific interactions with endocytic cargo proteins in mammals. Consistent with its role in mediating the endocytosis of cargos at the plasma membrane, μ2-YELLOW FLUORESCENT PROTEIN localized to transient foci at the plasma membrane, and loss of μ2 resulted in defects in bulk endocytosis. Furthermore, loss of μ2 led to increased accumulation of YELLOW FLUORESCENT PROTEIN-CESA6 particles at the plasma membrane. Our results suggest that CESA represents a new class of CME cargo proteins and that plant cells might regulate cellulose synthesis by controlling the abundance of active CESA complexes at the plasma membrane through CME.Cellulose microfibrils, as the major load-bearing polymers in plant cell walls, are the predominant component that enforces asymmetric cell expansion (Green, 1962). In higher plants, cellulose is synthesized by multimeric rosettes, which are also referred to as cellulose synthase complexes (CSCs; Kimura et al., 1999). Genetic and coimmunoprecipitation studies have indicated that CELLULOSE SYNTHASE1 (CESA1), CESA3, and CESA6-like (CESA6, CESA2, CESA5, and CESA9) isoforms are constituents of CSCs during primary cell wall synthesis (Persson et al., 2005; Desprez et al., 2007; Persson et al., 2007; Wang et al., 2008), whereas CESA4, CESA7, and CESA8 are implicated in the cellulose synthesis of secondary cell walls (Taylor et al., 1999, 2003; Brown et al., 2005). Knowledge about cellulose synthesis has recently been enhanced by the development of a system whereby the dynamics of CESA can be imaged in living cells (Paredez et al., 2006; Desprez et al., 2007). In agreement with earlier transmission electron microscopy studies in which rosettes were visualized in Golgi cisternae, vesicles, and at the plasma membrane (Haigler and Brown, 1986), fluorescent protein tagging of CESA has identified CESA localization at the plasma membrane, in Golgi bodies, and in small intracellular compartments (Paredez et al., 2006; Desprez et al., 2007; Crowell et al., 2009; Gutierrez et al., 2009; Gu et al., 2010; Lei et al., 2012; Li et al., 2012b).Assuming that cellulose synthesis occurs solely at the plasma membrane, the trafficking of CSCs to and from the plasma membrane may act as a significant regulatory mechanism. Although the mechanistic details of CESA trafficking are lacking, live cell imaging has shown that CESA localizes to various subcellular compartments. A subset of CESAs colocalize with markers of the trans-Golgi network (TGN)/early endosome (EE), an organelle that is part of both the secretory and endocytic pathways in Arabidopsis (Arabidopsis thaliana; Dettmer et al., 2006; Lam et al., 2007; Crowell et al., 2009, 2010; Viotti et al., 2010). CESAs also localize to microtubule-associated cellulose synthase compartments (MASCs) and small CESA-containing compartments (SmaCCs). The exact function of SmaCCs/MASCs is unknown, but it has been proposed that SmaCCs/MASCs might result from the internalization of CSCs or might act in the delivery of CSCs to the plasma membrane (Crowell et al., 2009, 2010; Gutierrez et al., 2009).Clathrin-mediated endocytosis (CME) has been shown to be a major endocytic pathway in Arabidopsis (Holstein, 2002; Samaj et al., 2005; Dhonukshe et al., 2007; Kleine-Vehn and Friml, 2008; Chen et al., 2011; Beck et al., 2012; Wang et al., 2013), although there is also evidence of clathrin-independent endocytosis mechanisms (Bandmann and Homann, 2012). The function of many CME proteins has been extensively characterized in mammals (McMahon and Boucrot, 2011), and homologs of many CME components are encoded by the Arabidopsis genome, including multiple copies of clathrin H chain and clathrin light chain (CLC), all four subunits of the heterotetrameric ADAPTOR PROTEIN COMPLEX2 (AP2) complex, dynamin-related proteins, and accessory proteins such as AP180 (Holstein, 2002; Chen et al., 2011); however, many CME components have yet to be characterized in plants.It has been suggested that CME might also function in controlling cell wall metabolism. For example, dividing and growing cells internalize cross-linked cell wall pectins, which might allow for cell wall remodeling (Baluska et al., 2002, 2005; Samaj et al., 2004). Moreover, the importance of endocytosis for cell wall morphogenesis is apparent from the functional characterization of proteins involved in CME. A dynamin-related protein, DRP1A, plays a significant role in endocytosis and colocalizes with CLC (Collings et al., 2008; Konopka and Bednarek, 2008). Defective endocytosis in RADIAL SWELLING9 (rsw9) plants, which contain a mutation in DRP1A, results in cellulose deficiency and defects in cell elongation (Collings et al., 2008). A mutation in rice, brittle culm3 (bc3), was mapped to the dynamin-related gene OsDRP2A, which has been proposed to function in CME. The brittle-culm phenotype in this mutant was attributed to cellulose deficiency (Xiong et al., 2010). Although the abundance of OsCESA4 was also altered in bc3, it remains unclear whether the cellulose deficiency of either bc3 or rsw9 results directly from perturbations in CESA trafficking.To identify proteins involved in the regulation of cellulose biosynthesis, a yeast two-hybrid (Y2H) screen was performed in which the central domain of CESA6 (CESA6CD) was used as bait to screen an Arabidopsis complementary DNA library for potential interaction partners of CESA6 (Gu et al., 2010; Gu and Somerville, 2010). The Y2H screen identified μ2 as a putative interaction partner of CESA6CD. The mammalian homolog of μ2 is the medium subunit of the tetrameric AP2, which acts as the core of the CME machinery by docking to the plasma membrane while concomitantly recruiting cargo proteins, clathrin triskelia, and accessory proteins to the sites of endocytosis (Jackson et al., 2010; McMahon and Boucrot, 2011; Cocucci et al., 2012). In this study, we provide evidence that μ2 plays a role in CME in Arabidopsis, that CESAs are a new set of CME cargo proteins, and that plant cells might regulate cellulose synthesis by controlling the abundance of CSCs at the plasma membrane through CME. To our knowledge, this study is the first to show the affect of an AP2 complex component on endocytosis in plants and the first to visualize an AP2 complex component in living plant cells. Furthermore, our data suggest that the role of AP2 in plants may differ from what has been shown in animals.     

On the Inside

Cellulose synthase complexes (CSCs) at the plasma membrane (PM) are aligned with cortical microtubules (MTs) and direct the biosynthesis of cellulose. The mechanism of the interaction between CSCs and MTs, and the cellular determinants that control the delivery of CSCs at the PM, are not yet well understood. We identified a unique small molecule, CESA TRAFFICKING INHIBITOR (CESTRIN), which reduces cellulose content and alters the anisotropic growth of Arabidopsis (Arabidopsis thaliana) hypocotyls. We monitored the distribution and mobility of fluorescently labeled cellulose synthases (CESAs) in live Arabidopsis cells under chemical exposure to characterize their subcellular effects. CESTRIN reduces the velocity of PM CSCs and causes their accumulation in the cell cortex. The CSC-associated proteins KORRIGAN1 (KOR1) and POM2/CELLULOSE SYNTHASE INTERACTIVE PROTEIN1 (CSI1) were differentially affected by CESTRIN treatment, indicating different forms of association with the PM CSCs. KOR1 accumulated in bodies similar to CESA; however, POM2/CSI1 dissociated into the cytoplasm. In addition, MT stability was altered without direct inhibition of MT polymerization, suggesting a feedback mechanism caused by cellulose interference. The selectivity of CESTRIN was assessed using a variety of subcellular markers for which no morphological effect was observed. The association of CESAs with vesicles decorated by the trans-Golgi network-localized protein SYNTAXIN OF PLANTS61 (SYP61) was increased under CESTRIN treatment, implicating SYP61 compartments in CESA trafficking. The properties of CESTRIN compared with known CESA inhibitors afford unique avenues to study and understand the mechanism under which PM-associated CSCs are maintained and interact with MTs and to dissect their trafficking routes in etiolated hypocotyls.Plant cell expansion and anisotropic cell growth are driven by vacuolar turgor pressure and cell wall extensibility, which in a dynamic and restrictive manner direct cell morphogenesis (Baskin, 2005). Cellulose is the major load-bearing component of the cell wall and is thus a major determinant for anisotropic growth (Baskin, 2001). Cellulose is made up of β-1,4-linked glucan chains that may aggregate to form microfibrils holding 18 to 36 chains (Somerville, 2006; Fernandes et al., 2011; Jarvis, 2013; Newman et al., 2013; Thomas et al., 2013). In contrast to cell wall structural polysaccharides, including pectin and hemicellulose, which are synthesized by Golgi-localized enzymes, cellulose is synthesized at the plasma membrane (PM) by cellulose synthase complexes (CSCs; Somerville, 2006; Scheller and Ulvskov, 2010; Atmodjo et al., 2013). The cellulose synthases (CESAs) are the principal catalytic units of cellulose biosynthesis and in higher plants are organized into globular rosettes (Haigler and Brown, 1986). For their biosynthetic function, each primary cell wall CSC requires a minimum of three catalytic CESA proteins (Desprez et al., 2007; Persson et al., 2007).On the basis of observations that cellulose microfibrils align with cortical microtubules (MTs) and that MT disruption leads to a loss of cell expansion, it was hypothesized that cortical MTs guide the deposition and, therefore, the orientation of cellulose (Green, 1962; Ledbetter and Porter, 1963; Baskin, 2001; Bichet et al., 2001; Sugimoto et al., 2003; Baskin et al., 2004; Wasteneys and Fujita, 2006). Confocal microscopy of CESA fluorescent fusions has advanced our understanding of CESA trafficking and dynamics. CSCs are visualized as small particles moving within the plane of the PM, with an average velocity of approximately 200 to 400 nm min⁻¹. Their movement in linear tracks along cortical MTs (Paredez et al., 2006) supports the MT-cellulose alignment hypothesis.Our current understanding of cellulose synthesis suggests that CESAs are assembled into CSCs in either the endoplasmic reticulum (ER) or the Golgi apparatus and trafficked by vesicles to the PM (Bashline et al., 2014; McFarlane et al., 2014). The presence of CESAs in isolated Golgi and vesicles from the trans-Golgi network (TGN) has been established by proteomic studies (Dunkley et al., 2006; Drakakaki et al., 2012; Nikolovski et al., 2012; Parsons et al., 2012; Groen et al., 2014). Their localization at the TGN has been corroborated by electron microscopy and colocalization with TGN markers, such as vacuolar H⁺-ATP synthase subunit a1 (VHA-a1), and the Soluble NSF Attachment Protein Receptor (SNARE) protein SYNTAXIN OF PLANTS41 (SYP41), SYP42, and SYP61 (Crowell et al., 2009; Gutierrez et al., 2009; Drakakaki et al., 2012). A population of post-Golgi compartments carrying CSCs, referred to as microtubule-associated cellulose synthase compartments (MASCs) or small cellulose synthase compartments (SmaCCs), may be associated with MTs or actin filaments and are thought to be directly involved in either CSC delivery to, or internalization from, the PM (Crowell et al., 2009; Gutierrez et al., 2009).In addition to the CESAs, auxiliary proteins have been identified that play a vital role in the cellulose-synthesizing machinery. These include COBRA (Roudier et al., 2005), the endoglucanase KORRIGAN1 (KOR1; Lane et al., 2001; Lei et al., 2014b; Vain et al., 2014), and the recently identified POM-POM2/CELLULOSE SYNTHASE INTERACTIVE PROTEIN1 (POM2/CSI1; Gu et al., 2010; Bringmann et al., 2012). The latter protein functions as a linker between the cortical MTs and CSCs, as genetic lesions in POM2/CSI1 result in a lower incidence of coalignment between CSCs and cortical MTs (Bringmann et al., 2012). Given the highly regulated process of cellulose biosynthesis and deposition, it can be expected that many more accessory proteins participate in the delivery of CSCs and their interaction with MTs. Identification of these unique CSC-associated proteins can ultimately provide clues for the mechanisms behind cell growth and cell shape formation.Arabidopsis (Arabidopsis thaliana) mutants with defects in the cellulose biosynthetic machinery exhibit a loss of anisotropic growth, which results in organ swelling. This phenotype may be used as a diagnostic tool in genetic screens to identify cellulose biosynthetic and CSC auxiliary proteins (Mutwil et al., 2008). Chemical inhibitors complement genetic lesions to perturb, study, and control the cellular and physiological function of proteins (Drakakaki et al., 2009). A plethora of bioactive small molecules have been identified, and their analytical use contributes to our understanding of cellulose biosynthesis and CESA subcellular behavior (for review, see Brabham and Debolt, 2012). Small molecule treatment can induce distinct characteristic subcellular CESA patterns that can be broadly grouped into three categories (Brabham and Debolt, 2012). The first is characterized by the depletion of CESAs from the PM and their accumulation in cytosolic compartments, as observed for the herbicide isoxaben {N-[3-(1-ethyl-1-methylpropyl)-5-isoxazolyl]-2,6-dimethyoxybenzamide}, CGA 325615 [1-cyclohexyl-5-(2,3,4,5,6-pentafluorophe-noxyl)-1λ4,2,4,6-thiatriazin-3-amine], thaxtomin A (4-nitroindol-3-yl containing 2,5-dioxopiperazine), AE F150944 [N2-(1-ethyl-3-phenylpropyl)-6-(1-fluoro-1-methylethyl)-1,3,5-triazine-2,4-di-amine], and quinoxyphen [4-(2-bromo-4,5-dimethoxyphenyl)-3,4-dihydro-¹H-benzo-quinolin-2-one]; (Paredez et al., 2006; Bischoff et al., 2009; Crowell et al., 2009; Gutierrez et al., 2009; Harris et al., 2012). The second displays hyperaccumulation of CESAs at the PM, as seen for the herbicides dichlobenil (2,6-dichlorobenzonitrile) and indaziflam {N-[(1R,2S)-2,3-dihydro-2,6-dimethyl-¹H-inden-1-yl)-6-(1-fluoroethyl]-1,3,5-triazine-2,4-diamine} (Herth, 1987; DeBolt et al., 2007b; Brabham et al., 2014). The third exhibits disturbance of both CESAs and MTs and alters CESA trajectories at the PM, as exemplified by morlin (7-ethoxy-4-methylchromen-2-one; DeBolt et al., 2007a). Unique compounds inducing a phenotype combining CESA accumulation in intermediate compartments and disruption of CSC-MT interactions can contribute to both the identification of the accessory proteins linking CSCs with MTs and the vesicular delivery mechanisms of CESAs.In this study, we identified and characterized a unique cellulose deposition inhibitor, the small molecule CESA TRAFFICKING INHIBITOR (CESTRIN), which affects the localization pattern of CSCs and their interacting proteins in a unique way. The induction of cytoplasmic CESTRIN bodies might provide further clues for trafficking routes that carry CESAs to the PM.     

Unidirectional Movement of Cellulose Synthase Complexes in Arabidopsis Seed Coat Epidermal Cells Deposit Cellulose Involved in Mucilage Extrusion,Adherence, and Ray Formation

CELLULOSE SYNTHASE5 (CESA5) synthesizes cellulose necessary for seed mucilage adherence to seed coat epidermal cells of Arabidopsis (Arabidopsis thaliana). The involvement of additional CESA proteins in this process and details concerning the manner in which cellulose is deposited in the mucilage pocket are unknown. Here, we show that both CESA3 and CESA10 are highly expressed in this cell type at the time of mucilage synthesis and localize to the plasma membrane adjacent to the mucilage pocket. The isoxaben resistant1-1 and isoxaben resistant1-2 mutants affecting CESA3 show defects consistent with altered mucilage cellulose biosynthesis. CESA3 can interact with CESA5 in vitro, and green fluorescent protein-tagged CESA5, CESA3, and CESA10 proteins move in a linear, unidirectional fashion around the cytoplasmic column of the cell, parallel with the surface of the seed, in a pattern similar to that of cortical microtubules. Consistent with this movement, cytological evidence suggests that the mucilage is coiled around the columella and unwinds during mucilage extrusion to form a linear ray. Mutations in CESA5 and CESA3 affect the speed of mucilage extrusion and mucilage adherence. These findings imply that cellulose fibrils are synthesized in an ordered helical array around the columella, providing a distinct structure to the mucilage that is important for both mucilage extrusion and adherence.The epidermal cells of Arabidopsis (Arabidopsis thaliana) seed coats produce two distinct secondary cell walls: pectin-rich mucilage and cellulose-rich columellae (Western et al., 2000). When seeds are hydrated, mucilage expands rapidly, rupturing the outer tangential cell wall and forming a mucilage capsule that surrounds the seed. Seed coat mucilage is composed primarily of rhamnogalacturonan I (RG I) and also contains homogalacturonan (HG), hemicelluloses (such as xylans and glucomannans), and cellulose (for review, see Haughn and Western, 2012). Extruded mucilage consists of an outer, nonadherent fraction and an inner, adherent fraction (Western et al., 2000, 2001; Macquet et al., 2007a). The adherent and nonadherent mucilage layers differ in the amount of methylesterified HG (Rautengarten et al., 2008; Saez-Aguayo et al., 2013; Voiniciuc et al., 2013), galactans (Dean et al., 2007; Macquet et al., 2007b), arabinans (Arsovski et al., 2009), mannans (Yu et al., 2014), and cellulose (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011), all of which influence the physical properties of the layers.Adherent mucilage has a distinct structure, which can be examined using cell wall dyes and antibodies. When treated with cellulose-specific dyes, densely stained rays extend from the top of each columella to the outer edge of the adherent layer, many cell lengths above the seed surface (Mendu et al., 2011; Sullivan et al., 2011). Cytological evidence indicates that cellulose, pectins, and mannans are components of the ray (Haughn and Western, 2012; Griffiths et al., 2014; North et al., 2014; Yu et al., 2014), although the exact manner in which they are assembled is unknown.Cellulose is abundant in mucilage rays and mediates adherence. Loss-of-function mutations in CELLULOSE SYNTHASE5 (CESA5) result in reduced cellulose levels and increased detachment of mucilage from the seed (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011; Griffiths et al., 2014). How a reduction in cellulose results in a loss of adherence is still unknown, but it likely involves interaction with other mucilage components such as pectin and arabinogalactan proteins (Griffiths et al., 2014). Since cesa5 mutants still have some cellulose in the rays of the adherent mucilage halo (Mendu et al., 2011; Sullivan et al., 2011), additional cellulose synthases must be involved in mucilage cellulose biosynthesis.The Arabidopsis genome encodes 10 different CESAs (Delmer, 1999; Richmond and Somerville, 2000). Multiple lines of evidence suggest that three different CESAs are required to form one active cellulose synthase complex (CSC; for review, see Somerville, 2006). CSCs are membrane-bound protein complexes that synthesize cellulose microfibrils in the apoplast (for review, see Somerville, 2006; Endler and Persson, 2011; Lei et al., 2012). CESA1, CESA3, and CESA6 are considered the core components of the primary wall CSC (Desprez et al., 2007; Persson et al., 2007). CESA2, CESA5, and CESA9 are partially redundant to CESA6 in primary wall biosynthesis, and genetic evidence suggests that each of these CESA polypeptides can form a functional CSC with CESA3 and CESA1 (Desprez et al., 2007; Persson et al., 2007). CESA10 is expressed in young plants, stems, floral tissue, and the base of rosette leaves (Beeckman et al., 2002; Doblin et al., 2002), but its function in cellulose biosynthesis is unclear. Other cesa mutant lines have been examined for altered mucilage phenotypes (cesa1, radially swollen1 [Burn et al., 2002; Sullivan et al., 2011], cesa2, cesa6, and cesa9 [Mendu et al., 2011]; CESA3, je5 [Sullivan et al., 2011] and cesa10-1 [Sullivan et al., 2011]); to date, only CESA5 has been shown to be required for cellulose biosynthesis during mucilage deposition.Two mutant alleles of CESA3, isoxaben resistant1-1 (ixr1-1) and ixr1-2, were isolated in a screen for resistance to the herbicide isoxaben (Scheible et al., 2001). Isoxaben inhibits the incorporation of Glc into the emerging cellulose polymer and is considered a potent and specific inhibitor of cellulose biosynthesis (Heim et al., 1990). Homozygous ixr1-1 and ixr1-2 lines show increased resistance to the herbicide, and the mutations causing this resistance were mapped to the genomic locus of CESA3 (Heim et al., 1990; Scheible et al., 2001). The ixr1-1 and ixr1-2 mutations cause amino acid substitutions near the C terminus of the CESA3 protein. ixr1-1 causes a Gly-to-Asn substitution (G998A) located in a transmembrane domain, while ixr1-2 contains a Thr-to-Ile substitution (T942I) in an apoplastic region of the protein between two transmembrane domains (Scheible et al., 2001). Recently, the ixr1-2 allele was shown to affect the velocity of CSCs in the plasma membrane, which consequently modifies cellulose crystallinity in the cell wall (Harris et al., 2012). It is not exactly clear how the ixr1-1 mutation affects cellulose biosynthesis. The effects of either of these mutations on seed coat mucilage have not been investigated.Since mucilage is composed primarily of pectins with smaller amounts of cellulose, seed coat epidermal cells represent an excellent system to study cellulose biosynthesis and interactions between cellulose and other wall components in muro. In this study, we investigated how cellulose is synthesized and deposited in seed coat epidermal cells. We show that at least three different CESA proteins are highly expressed in the seed coat epidermis during mucilage biosynthesis. These CESAs are oriented and move in a linear fashion around the cytoplasmic column of each cell in an identical pattern to cortical microtubules. In addition, we provide evidence that the adherent mucilage has a helical structure that expands and unwinds during extrusion to form the mucilage ray. We propose that during seed coat epidermal cell development, the biosynthesis of cellulose predetermines the structure of rays in the adherent mucilage layer.     

Xyloglucan Deficiency Disrupts Microtubule Stability and Cellulose Biosynthesis in Arabidopsis,Altering Cell Growth and Morphogenesis

Xyloglucan constitutes most of the hemicellulose in eudicot primary cell walls and functions in cell wall structure and mechanics. Although Arabidopsis (Arabidopsis thaliana) xxt1 xxt2 mutants lacking detectable xyloglucan are viable, they display growth defects that are suggestive of alterations in wall integrity. To probe the mechanisms underlying these defects, we analyzed cellulose arrangement, microtubule patterning and dynamics, microtubule- and wall-integrity-related gene expression, and cellulose biosynthesis in xxt1 xxt2 plants. We found that cellulose is highly aligned in xxt1 xxt2 cell walls, that its three-dimensional distribution is altered, and that microtubule patterning and stability are aberrant in etiolated xxt1 xxt2 hypocotyls. We also found that the expression levels of microtubule-associated genes, such as MAP70-5 and CLASP, and receptor genes, such as HERK1 and WAK1, were changed in xxt1 xxt2 plants and that cellulose synthase motility is reduced in xxt1 xxt2 cells, corresponding with a reduction in cellulose content. Our results indicate that loss of xyloglucan affects both the stability of the microtubule cytoskeleton and the production and patterning of cellulose in primary cell walls. These findings establish, to our knowledge, new links between wall integrity, cytoskeletal dynamics, and wall synthesis in the regulation of plant morphogenesis.The primary walls of growing plant cells are largely constructed of cellulose and noncellulosic matrix polysaccharides that include hemicelluloses and pectins (Carpita and Gibeaut, 1993; Somerville et al., 2004; Cosgrove, 2005). Xyloglucan (XyG) is the most abundant hemicellulose in the primary walls of eudicots and is composed of a β-1,4-glucan backbone with side chains containing Xyl, Gal, and Fuc (Park and Cosgrove, 2015). XyG is synthesized in the Golgi apparatus before being secreted to the apoplast, and its biosynthesis requires several glycosyltransferases, including β-1,4-glucosyltransferase, α-1,6-xylosyltransferase, β-1,2-galactosyltransferase, and α-1,2-fucosyltransferase activities (Zabotina, 2012). Arabidopsis (Arabidopsis thaliana) XYLOGLUCAN XYLOSYLTRANSFERASE1 (XXT1) and XXT2 display xylosyltransferase activity in vitro (Faik et al., 2002; Cavalier and Keegstra, 2006), and strikingly, no XyG is detectable in the walls of xxt1 xxt2 double mutants (Cavalier et al., 2008; Park and Cosgrove, 2012a), suggesting that the activity of XXT1 and XXT2 are required for XyG synthesis, delivery, and/or stability.Much attention has been paid to the interactions between cellulose and XyG over the past 40 years. Currently, there are several hypotheses concerning the nature of these interactions (Park and Cosgrove, 2015). One possibility is that XyGs bind directly to cellulose microfibrils (CMFs). Recent data indicating that crystalline cellulose cores are surrounded with hemicelluloses support this hypothesis (Dick-Pérez et al., 2011). It is also possible that XyG acts as a spacer-molecule to prevent CMFs from aggregating in cell walls (Anderson et al., 2010) or as an adapter to link cellulose with other cell wall components, such as pectin (Cosgrove, 2005; Cavalier et al., 2008). XyG can be covalently linked to pectin (Thompson and Fry, 2000; Popper and Fry, 2005, 2008), and NMR data demonstrate that pectins and cellulose might interact to a greater extent than XyG and cellulose in native walls (Dick-Pérez et al., 2011). Alternative models exist for how XyG-cellulose interactions influence primary wall architecture and mechanics. One such model posits that XyG chains act as load-bearing tethers that bind to CMFs in primary cell walls to form a cellulose-XyG network (Carpita and Gibeaut, 1993; Pauly et al., 1999; Somerville et al., 2004; Cosgrove, 2005). However, results have been accumulating against this tethered network model, leading to an alternative model in which CMFs make direct contact, in some cases mediated by a monolayer of xyloglucan, at limited cell wall sites dubbed “biomechanical hotspots,” which are envisioned as the key sites of cell wall loosening during cell growth (Park and Cosgrove, 2012a; Wang et al., 2013; Park and Cosgrove, 2015). Further molecular, biochemical, and microscopy experiments are required to help distinguish which aspects of the load-bearing, spacer/plasticizer, and/or hotspot models most accurately describe the functions of XyG in primary walls.Cortical microtubules (MTs) direct CMF deposition by guiding cellulose synthase complexes in the plasma membrane (Baskin et al., 2004; Paredez et al., 2006; Emons et al., 2007; Sánchez-Rodriguez et al., 2012), and the patterned deposition of cellulose in the wall in turn can help determine plant cell anisotropic growth and morphogenesis (Baskin, 2005). Disruption of cortical MTs by oryzalin, a MT-depolymerizing drug, alters the alignment of CMFs, suggesting that MTs contribute to CMF organization (Baskin et al., 2004). CELLULOSE SYNTHASE (CESA) genes, including CESA1, CESA3, and CESA6, are required for normal CMF synthesis in primary cell walls (Kohorn et al., 2006; Desprez et al., 2007), and accessory proteins such as COBRA function in cellulose production (Lally et al., 2001). Live-cell imaging from double-labeled YFP-CESA6; CFP-ALPHA-1 TUBULIN (TUA1) Arabidopsis seedlings provides direct evidence that cortical MTs determine the trajectories of cellulose synthesis complexes (CSCs) and patterns of cellulose deposition (Paredez et al., 2006). Additionally, MT organization affects the rotation of cellulose synthase trajectories in the epidermal cells of Arabidopsis hypocotyls (Chan et al., 2010). Recently, additional evidence for direct guidance of CSCs by MTs has been provided by the identification of CSI1/POM2, which binds to both MTs and CESAs (Bringmann et al., 2012; Li et al., 2012). MICROTUBULE ORGANIZATION1 (MOR1) is essential for cortical MT organization (Whittington et al., 2001), but disruption of cortical MTs in the mor1 mutant does not greatly affect CMF organization (Sugimoto et al., 2003), and oryzalin treatment does not abolish CSC motility (Paredez et al., 2006).Conversely, the organization of cortical MTs can be affected by cellulose synthesis. Treatment with isoxaben, a cellulose synthesis inhibitor, results in disorganized cortical MTs in tobacco cells, suggesting that inhibition of cellulose synthesis affects MT organization (Fisher and Cyr, 1998), and treatment with 2,6-dichlorobenzonitrile, another cellulose synthesis inhibitor, alters MT organization in mor1 plants (Himmelspach et al., 2003). Cortical MT orientation in Arabidopsis roots is also altered in two cellulose synthesis-deficient mutants, CESA6^52-isx and kor1-3, suggesting that CSC activity can affect MT arrays (Paredez et al., 2008). Together, these results point to a bidirectional relationship between cellulose synthesis/patterning and MT organization.MTs influence plant organ morphology, but the detailed mechanisms by which they do so are incompletely understood. The dynamics and stability of cortical MTs are also affected by MT-associated proteins (MAPs). MAP18 is a MT destabilizing protein that depolymerizes MTs (Wang et al., 2007), MAP65-1 functions as a MT crosslinker, and MAP70-1 functions in MT assembly (Korolev et al., 2005; Lucas et al., 2011). MAP70-5 stabilizes existing MTs to maintain their length, and its overexpression induces right-handed helical growth (Korolev et al., 2007); likewise, MAP20 overexpression results in helical cell twisting (Rajangam et al., 2008). CLASP promotes microtubule stability, and its mutant is hypersensitive to microtubule-destabilizing drug oryzalin (Ambrose et al., 2007). KATANIN1 (KTN1) is a MT-severing protein that can sever MTs into short fragments and promote the formation of thick MT bundles that ultimately depolymerize (Stoppin-Mellet et al., 2006), and loss of KTN1 function results in reduced responses to mechanical stress (Uyttewaal et al., 2012). In general, cortical MT orientation responds to mechanical signals and can be altered by applying force directly to the shoot apical meristem (Hamant et al., 2008). The application of external mechanical pressure to Arabidopsis leaves also triggers MT bundling (Jacques et al., 2013). Kinesins, including KINESIN-13A (KIN-13A) and FRAGILE FIBER1 (FRA1), have been implicated in cell wall synthesis (Cheung and Wu, 2011; Fujikura et al., 2014). The identification of cell wall receptors and sensors is beginning to reveal how plant cell walls sense and respond to external signals (Humphrey et al., 2007; Ringli, 2010); some of them, such as FEI1, FEI2, THESEUS1 (THE1), FERONIA (FER), HERCULES RECEPTOR KINASE1 (HERK1), WALL ASSOCIATED KINASE1 (WAK1), WAK2, and WAK4, have been characterized (Lally et al., 2001; Decreux and Messiaen, 2005; Kohorn et al., 2006; Xu et al., 2008; Guo et al., 2009; Cheung and Wu, 2011). However, the relationships between wall integrity, cytoskeletal dynamics, and wall synthesis have not yet been fully elucidated.In this study, we analyzed CMF patterning, MT patterning and dynamics, and cellulose biosynthesis in the Arabidopsis xxt1 xxt2 double mutant that lacks detectable XyG and displays altered growth (Cavalier et al., 2008; Park and Cosgrove, 2012a). To investigate whether and how XyG deficiency affects the organization of CMFs and cortical MTs, we observed CMF patterning in xxt1 xxt2 mutants and Col (wild-type) controls using atomic force microscopy (AFM), field emission scanning electron microscopy (FESEM), transmission electron microscopy (TEM), and confocal microscopy (Hodick and Kutschera, 1992; Derbyshire et al., 2007; Anderson et al., 2010; Zhang et al., 2014). We also generated transgenic Col and xxt1 xxt2 lines expressing GFP-MAP4 (Marc et al., 1998) and GFP-CESA3 (Desprez et al., 2007), and analyzed MT arrays and cellulose synthesis using live-cell imaging. Our results show that the organization of CMFs is altered, that MTs in xxt1 xxt2 mutants are aberrantly organized and are more sensitive to external mechanical pressure and the MT-depolymerizing drug oryzalin, and that cellulose synthase motility and cellulose content are decreased in xxt1 xxt2 mutants. Furthermore, real-time quantitative RT-PCR measurements indicate that the enhanced sensitivity of cortical MTs to mechanical stress and oryzalin in xxt1 xxt2 plants might be due to altered expression of MT-stabilizing and wall receptor genes. Together, these data provide insights into the connections between the functions of XyG in wall assembly, the mechanical integrity of the cell wall, cytoskeleton-mediated cellular responses to deficiencies in wall biosynthesis, and cell and tissue morphogenesis.     

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 Arabidopsis Endosomal Sorting Complex Required for Transport III Regulates Internal Vesicle Formation of the Prevacuolar Compartment and Is Required for Plant Development

We have established an efficient transient expression system with several vacuolar reporters to study the roles of endosomal sorting complex required for transport (ESCRT)-III subunits in regulating the formation of intraluminal vesicles of prevacuolar compartments (PVCs)/multivesicular bodies (MVBs) in plant cells. By measuring the distributions of reporters on/within the membrane of PVC/MVB or tonoplast, we have identified dominant negative mutants of ESCRT-III subunits that affect membrane protein degradation from both secretory and endocytic pathways. In addition, induced expression of these mutants resulted in reduction in luminal vesicles of PVC/MVB, along with increased detection of membrane-attaching vesicles inside the PVC/MVB. Transgenic Arabidopsis (Arabidopsis thaliana) plants with induced expression of ESCRT-III dominant negative mutants also displayed severe cotyledon developmental defects with reduced cell size, loss of the central vacuole, and abnormal chloroplast development in mesophyll cells, pointing out an essential role of the ESCRT-III complex in postembryonic development in plants. Finally, membrane dissociation of ESCRT-III components is important for their biological functions and is regulated by direct interaction among Vacuolar Protein Sorting-Associated Protein20-1 (VPS20.1), Sucrose Nonfermenting7-1, VPS2.1, and the adenosine triphosphatase VPS4/SUPPRESSOR OF K⁺ TRANSPORT GROWTH DEFECT1.Endomembrane trafficking in plant cells is complicated such that secretory, endocytic, and recycling pathways are usually integrated with each other at the post-Golgi compartments, among which, the trans-Golgi network (TGN) and prevacuolar compartment (PVC)/multivesicular body (MVB) are best studied (Tse et al., 2004; Lam et al., 2007a, 2007b; Müller et al., 2007; Foresti and Denecke, 2008; Hwang, 2008; Otegui and Spitzer, 2008; Robinson et al., 2008; Richter et al., 2009; Ding et al., 2012; Gao et al., 2014). Following the endocytic trafficking of a lipophilic dye, FM4-64, the TGN and PVC/MVB are sequentially labeled and thus are defined as the early and late endosome, respectively, in plant cells (Lam et al., 2007a; Chow et al., 2008). While the TGN is a tubular vesicular-like structure that may include several different microdomains and fit its biological function as a sorting station (Chow et al., 2008; Kang et al., 2011), the PVC/MVB is 200 to 500 nm in size with multiple luminal vesicles of approximately 40 nm (Tse et al., 2004). Membrane cargoes destined for degradation are sequestered into these tiny luminal vesicles and delivered to the lumen of the lytic vacuole (LV) via direct fusion between the PVC/MVB and the LV (Spitzer et al., 2009; Viotti et al., 2010; Cai et al., 2012). Therefore, the PVC/MVB functions between the TGN and LV as an intermediate organelle and decides the fate of membrane cargoes in the LV.In yeast (Saccharomyces cerevisiae), carboxypeptidase S (CPS) is synthesized as a type II integral membrane protein and sorted from the Golgi to the lumen of the vacuole (Spormann et al., 1992). Genetic analyses on the trafficking of CPS have led to the identification of approximately 17 class E genes (Piper et al., 1995; Babst et al., 1997, 2002a, 2002b; Odorizzi et al., 1998; Katzmann et al., 2001) that constitute the core endosomal sorting complex required for transport (ESCRT) machinery. The evolutionarily conserved ESCRT complex consists of several functionally different subcomplexes, ESCRT-0, ESCRT-I, ESCRT-II, and ESCRT-III and the ESCRT-III-associated/Vacuolar Protein Sorting4 (VPS4) complex. Together, they form a complex protein-protein interaction network that coordinates sorting of cargoes and inward budding of the membrane on the MVB (Hurley and Hanson, 2010; Henne et al., 2011). Cargo proteins carrying ubiquitin signals are thought to be passed from one ESCRT subcomplex to the next, starting with their recognition by ESCRT-0 (Bilodeau et al., 2002, 2003; Hislop and von Zastrow, 2011; Le Bras et al., 2011; Shields and Piper, 2011; Urbé, 2011). ESCRT-0 recruits the ESCRT-I complex, a heterotetramer of VPS23, VPS28, VPS37, and MVB12, from the cytosol to the endosomal membrane (Katzmann et al., 2001, 2003). The C terminus of VPS28 interacts with the N terminus of VPS36, a member of the ESCRT-II complex (Kostelansky et al., 2006; Teo et al., 2006). Then, cargoes passed from ESCRT-I and ESCRT-II are concentrated in certain membrane domains of the endosome by ESCRT-III, which includes four coiled-coil proteins and is sufficient to induce the membrane invagination (Babst et al., 2002b; Saksena et al., 2009; Wollert et al., 2009). Finally, the ESCRT components are disassociated from the membrane by the adenosine triphosphatase (ATPase) associated with diverse cellular activities (AAA) VPS4/SUPPRESSOR OF K+ TRANSPORT GROWTH DEFECT1 (SKD1) before releasing the internal vesicles (Babst et al., 1997, 1998).Putative homologs of ESCRT-I–ESCRT-III and ESCRT-III-associated components have been identified in plants, except for ESCRT-0, which is only present in Opisthokonta (Winter and Hauser, 2006; Leung et al., 2008; Schellmann and Pimpl, 2009). To date, only a few plant ESCRT components have been studied in detail. The Arabidopsis (Arabidopsis thaliana) AAA ATPase SKD1 localized to the PVC/MVB and showed ATPase activity that was regulated by Lysosomal Trafficking Regulator-Interacting Protein5, a plant homolog of Vps Twenty Associated1 Protein (Haas et al., 2007). Expression of the dominant negative form of SKD1 caused an increase in the size of the MVB and a reduction in the number of internal vesicles (Haas et al., 2007). This protein also contributes to the maintenance of the central vacuole and might be associated with cell cycle regulation, as leaf trichomes expressing its dominant negative mutant form lost the central vacuole and frequently contained multiple nuclei (Shahriari et al., 2010). Double null mutants of CHARGED MULTIVESICULAR BODY PROTEIN, chmp1achmp1b, displayed severe growth defects and were seedling lethal. This may be due to the mislocalization of plasma membrane (PM) proteins, including those involved in auxin transport such as PINFORMED1, PINFORMED2, and AUXIN-RESISTANT1, from the vacuolar degradation pathway to the tonoplast of the LV (Spitzer et al., 2009).Plant ESCRT components usually contain several homologs, with the possibility of functional redundancy. Single mutants of individual ESCRT components may not result in an obvious phenotype, whereas knockout of all homologs of an ESCRT component by generating double or triple mutants may be lethal to the plant. As a first step to carry out systematic analysis on each ESCRT complex in plant cells, here, we established an efficient analysis system to monitor the localization changes of four vacuolar reporters that accumulate either in the lumen (LRR84A-GFP, EMP12-GFP, and aleurain-GFP) or on the tonoplast (GFP-VIT1) of the LV and identified several ESCRT-III dominant negative mutants. We reported that ESCRT-III subunits were involved in the release of PVC/MVB’s internal vesicles from the limiting membrane and were required for membrane protein degradation from secretory and endocytic pathways. In addition, transgenic Arabidopsis plants with induced expression of ESCRT-III dominant negative mutants showed severe cotyledon developmental defects. We also showed that membrane dissociation of ESCRT-III subunits was regulated by direct interaction with SKD1.