MUCILAGE-RELATED10 Produces Galactoglucomannan That Maintains Pectin and Cellulose Architecture in Arabidopsis Seed Mucilage

Plants invest a lot of their resources into the production of an extracellular matrix built of polysaccharides. While the composition of the cell wall is relatively well characterized, the functions of the individual polymers and the enzymes that catalyze their biosynthesis remain poorly understood. We exploited the Arabidopsis (Arabidopsis thaliana) seed coat epidermis (SCE) to study cell wall synthesis. SCE cells produce mucilage, a specialized secondary wall that is rich in pectin, at a precise stage of development. A coexpression search for MUCILAGE-RELATED (MUCI) genes identified MUCI10 as a key determinant of mucilage properties. MUCI10 is closely related to a fenugreek (Trigonella foenumgraecum) enzyme that has in vitro galactomannan α-1,6-galactosyltransferase activity. Our detailed analysis of the muci10 mutants demonstrates that mucilage contains highly branched galactoglucomannan (GGM) rather than unbranched glucomannan. MUCI10 likely decorates glucomannan, synthesized by CELLULOSE SYNTHASE-LIKE A2, with galactose residues in vivo. The degree of galactosylation is essential for the synthesis of the GGM backbone, the structure of cellulose, mucilage density, as well as the adherence of pectin. We propose that GGM scaffolds control mucilage architecture along with cellulosic rays and show that Arabidopsis SCE cells represent an excellent model in which to study the synthesis and function of GGM. Arabidopsis natural varieties with defects similar to muci10 mutants may reveal additional genes involved in GGM synthesis. Since GGM is the most abundant hemicellulose in the secondary walls of gymnosperms, understanding its biosynthesis may facilitate improvements in the production of valuable commodities from softwoods.The plant cell wall is the key determinant of plant growth (Cosgrove, 2005) and represents the most abundant source of biopolymers on the planet (Pauly and Keegstra, 2010). Consequently, plants invest a lot of their resources into the production of this extracellular structure. Thus, it is not surprising that approximately 15% of Arabidopsis (Arabidopsis thaliana) genes are likely dedicated to the biosynthesis and modification of cell wall polymers (Carpita et al., 2001). Plant walls consist mainly of polysaccharides (cellulose, hemicellulose, and pectin) but also contain lignin and glycoproteins. While the biochemical structure of each wall component has been relatively well characterized, the molecular players involved in their biogenesis remain poorly understood (Keegstra, 2010). The functions of the individual polymers, and how they are assembled into a three-dimensional matrix, are also largely unknown (Burton et al., 2010; Burton and Fincher, 2012).Significant breakthroughs in cell wall research have been achieved through the examination of specialized plant tissues that contain elevated levels of a single polysaccharide (Pauly and Keegstra, 2010). Some species, particularly legumes, accumulate large amounts of the hemicellulose galactomannan during secondary wall thickening of the seed (Srivastava and Kapoor, 2005). Analysis of the developing fenugreek (Trigonella foenumgraecum) endosperm led to the purification of a GALACTOMANNAN GALACTOSYLTRANSFERASE (TfGMGT), the first glycosyltransferase (GT) whose activity in plant cell wall synthesis was demonstrated in vitro (Scheller and Ulvskov, 2010). TfGMGT catalyzes the decoration of mannan chains with single α-1,6-galactosyl residues (Edwards et al., 1999). A similar approach in guar (Cyamopsis tetragonoloba) seeds revealed that the β-1,4-linked mannan backbone is synthesized by a member of the CELLULOSE SYNTHASE-LIKE A (CSLA) protein family (Dhugga et al., 2004).Galactomannan functions as a storage polymer in the endosperm of the aforementioned seeds, analogous to starch in cereal grains (Dhugga et al., 2004), but it also has important rheological properties in the cell wall that have been exploited to produce valuable stabilizers and gelling agents for human consumption (Srivastava and Kapoor, 2005). The Man-to-Gal ratio is essential for the application of galactomannan gums in the food industry (Edwards et al., 1992). This is because unsubstituted mannan chains can interact via hydrogen bonds to produce crystalline microfibrils similar to cellulose (Millane and Hendrixson, 1994). Indeed, some algae that lack cellulose employ mannan fibrils as a structural material (Preston, 1968). The addition of Gal branches to the smooth, ribbon-like mannan chains creates hairy regions that limit self-association and promote gelation (Dea et al., 1977). All mannans are likely synthesized as highly substituted polymers that are trimmed in the cell wall (Scheller and Ulvskov, 2010).Generally, polysaccharides containing backbones of β-1,4-linked Man units can be classified as heteromannan (HM). Galactoglucomannan (GGM) is the main hemicellulose in gymnosperm secondary walls and, in contrast to galactomannan, has a backbone that contains both Glc and Man units (Pauly et al., 2013). HM is detected in most Arabidopsis cell types (Handford et al., 2003) and facilitates embryogenesis (Goubet et al., 2009), germination (Rodríguez-Gacio et al., 2012), tip growth (Bernal et al., 2008), and vascular development (Benová-Kákosová et al., 2006; Yin et al., 2011). In the last 10 years, in vitro mannan synthase activity has been demonstrated for recombinant CSLA proteins from many land plants (Liepman et al., 2005, 2007; Suzuki et al., 2006; Gille et al., 2011; Wang et al., 2012). HM synthesis may also involve CELLULOSE SYNTHASE-LIKE D (CSLD) enzymes and MANNAN SYNTHESIS-RELATED (MSR) accessory proteins (Yin et al., 2011; Wang et al., 2013), but their precise roles in relation to the CSLAs have not been established. Arabidopsis CSLA2, like most other isoforms, can use both GDP-Man and GDP-Glc as substrates in vitro (Liepman et al., 2005, 2007) and is responsible for stem glucomannan synthesis in vivo along with CSLA3 and CSLA7 (Goubet et al., 2009). CSLA2 also participates in the synthesis of glucomannan present in mucilage produced by seed coat epidermal (SCE) cells (Yu et al., 2014).Arabidopsis SCE cells represent an excellent genetic model in which to study the synthesis, polar secretion, and modification of polysaccharides, since these processes dominate a precise stage of seed coat development but are not essential for seed viability in laboratory conditions (Haughn and Western, 2012; North et al., 2014; Voiniciuc et al., 2015). Hydration of mature seeds in water releases a large gelatinous capsule, rich in the pectic polymer rhamnogalacturonan I, which can be easily stained or extracted (Macquet et al., 2007). Biochemical and cytological experiments indicate that Arabidopsis seed mucilage is more than just pectin and, in addition to cellulose, is likely to contain glycoproteins and at least two hemicellulosic polymers (Voiniciuc et al., 2015). There is mounting evidence that, despite their low abundance, these components play critical functions in seed mucilage architecture. The structure of homogalacturonan (HG), the major pectin in primary cell walls but a minor mucilage component, appears to be a key determinant of gelling properties and mucilage extrusion (Rautengarten et al., 2008; Saez-Aguayo et al., 2013; Voiniciuc et al., 2013). Mucilage attachment to seeds is maintained by the SALT OVERLY SENSITIVE5 glycoprotein and cellulose synthesized by multiple CELLULOSE SYNTHASE (CESA) isoforms (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011; Griffiths et al., 2014, 2015). From more than 35 genes that are reported to affect Arabidopsis seed mucilage properties (Voiniciuc et al., 2015), only CSLA2, CESA3, CESA5, GALACTURONOSYLTRANSFERASE11 (GAUT11; Caffall et al., 2009), and GAUT-LIKE5 (GATL5; Kong et al., 2013) are predicted to encode GTs. This highlights that, despite many detailed studies about mucilage production in SCE cells, the synthesis of its components remains poorly understood.To address this issue, we conducted a reverse genetic search for MUCILAGE-RELATED (MUCI) genes that may be required for polysaccharide biosynthesis. One of these, MUCI10, encodes a member of the Carbohydrate Active Enzymes family, GT34 (Lombard et al., 2014), which includes at least two enzymatic activities and seven Arabidopsis proteins (Keegstra and Cavalier, 2010). Five of them function as XYLOGLUCAN XYLOSYLTRANSFERASES (XXT1–XXT5) in vivo and/or in vitro (Faik et al., 2002; Cavalier et al., 2008; Vuttipongchaikij et al., 2012). MUCI10/GT7 (At2g22900) and its paralog GT6 (At4g37690) do not function as XXTs (Vuttipongchaikij et al., 2012) and are more closely related to the TfGMGT enzyme (Faik et al., 2002; Keegstra and Cavalier, 2010). MUCI10, also called GALACTOSYLTRANSFERASE-LIKE6 (GTL6), served as a Golgi marker in multiple proteomic studies of Arabidopsis callus cultures (Dunkley et al., 2004, 2006; Nikolovski et al., 2012, 2014). Nevertheless, the role of TfGMGT orthologs in Arabidopsis remained unknown. We show that MUCI10 is responsible for the extensive galactosylation of glucomannan in mucilage and influences glucomannan backbone synthesis, cellulose structure, and the distribution of pectin.     

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.     

SALT-OVERLY SENSITIVE5 Mediates Arabidopsis Seed Coat Mucilage Adherence and Organization through Pectins

Interactions between cell wall polymers are critical for establishing cell wall integrity and cell-cell adhesion. Here, we exploit the Arabidopsis (Arabidopsis thaliana) seed coat mucilage system to examine cell wall polymer interactions. On hydration, seeds release an adherent mucilage layer strongly attached to the seed in addition to a nonadherent layer that can be removed by gentle agitation. Rhamnogalacturonan I (RG I) is the primary component of adherent mucilage, with homogalacturonan, cellulose, and xyloglucan constituting minor components. Adherent mucilage contains rays composed of cellulose and pectin that extend above the center of each epidermal cell. CELLULOSE SYNTHASE5 (CESA5) and the arabinogalactan protein SALT-OVERLY SENSITIVE5 (SOS5) are required for mucilage adherence through unknown mechanisms. SOS5 has been suggested to mediate adherence by influencing cellulose biosynthesis. We, therefore, investigated the relationship between SOS5 and CESA5. cesa5-1 seeds show reduced cellulose, RG I, and ray size in adherent mucilage. In contrast, sos5-2 seeds have wild-type levels of cellulose but completely lack adherent RG I and rays. Thus, relative to each other, cesa5-1 has a greater effect on cellulose, whereas sos5-2 mainly affects pectin. The double mutant cesa5-1 sos5-2 has a much more severe loss of mucilage adherence, suggesting that SOS5 and CESA5 function independently. Double-mutant analyses with mutations in MUCILAGE MODIFIED2 and FLYING SAUCER1 that reduce mucilage release through pectin modification suggest that only SOS5 influences pectin-mediated adherence. Together, these findings suggest that SOS5 mediates adherence through pectins and does so independently of but in concert with cellulose synthesized by CESA5.Cellulosic cell walls are a defining feature of land plants. Primary cell walls are composed of three major classes of polysaccharides: cellulose, hemicelluloses, and pectins. In addition, approximately 10% of the primary cell wall is composed of protein (Burton et al., 2010). Cell walls provide mechanical support for the cell, and cell wall carbohydrates in the middle lamellae mediate cell-cell adhesion (Caffall and Mohnen, 2009). Current models of cell wall structure depict a cellulose-hemicellulose network embedded in an independent pectin gel (for review, see Albersheim et al., 2011). These components are believed to interact through both covalent and noncovalent bonds to provide structure and strength to the cell wall, although the relative importance of pectin and its interactions with the hemicellulose-cellulose network remain unclear (for review, see Cosgrove, 2005).Another gap in our understanding of cell wall structure and assembly is the role of arabinogalactan proteins (AGPs). AGPs are a family of evolutionarily conserved secreted proteins highly glycosylated with type II arabinogalactans, and they can be localized to the plasma membrane by a C-terminal glycophosphatidylinositol (GPI) lipid anchor (for review, see Schultz et al., 2000; Showalter, 2001; Johnson et al., 2003; Seifert and Roberts, 2007; Ellis et al., 2010). AGPs can be extensively modified in the cell wall; many glycosyl hydrolases can affect AGP function by cleaving their glycosyl side chains (Sekimata et al., 1989; Cheung et al., 1995; Wu et al., 1995; Kotake et al., 2005). The GPI anchor can also be cleaved, releasing the AGPs from the membrane into the cell wall (Schultz et al., 2000). Although their exact roles are still unclear, AGPs have been proposed to interact with cell wall polysaccharides, initiate intracellular signaling cascades, and influence a wide variety of biological processes (for review, see Seifert and Roberts, 2007; Ellis et al., 2010; Tan et al., 2013).Many fasciclin-like AGPs (FLAs), which contain at least one fasciclin domain (FAS) associated with protein-protein interactions, have been suggested to influence cellulose biosynthesis or organization (Seifert and Roberts, 2007; Li et al., 2010; MacMillan et al., 2010). FLA3 RNA interference lines have reduced intine cell wall biosynthesis and loss of Calcofluor white (a fluorescent dye specific for glycan molecules) staining in aborted pollen grains (Li et al., 2010). A fla11 fla12 double mutant was shown to have reduced cellulose deposition, altered cellulose microfibril angle, and reduced cell wall integrity (MacMillan et al., 2010). The fla11 fla12 double mutant also had reductions in arabinans, galactans, and rhamnose (MacMillan et al., 2010). FLA4/SALT-OVERLY SENSITIVE5 (SOS5) was identified in a screen for salt sensitivity in roots. The SOS5 gene encodes an FLA protein with a GPI anchor, two AGP-like domains, and two FAS domains (Shi et al., 2003). Plants homozygous for the loss-of-function conditional allele sos5-1 have thinner root cell walls that appear less organized (Shi et al., 2003). The presence of the two FAS domains has led to the suggestion that SOS5 may interact with other proteins, forming a network that strengthens the cell wall (Shi et al., 2003). SOS5 is involved in regulation of cell wall rheology through a pathway involving two Leu-rich repeat receptor-like kinases, FEI1 and FEI2 (Xu et al., 2008). SOS5 and FEI2 are also required for normal seed coat mucilage adherence and hypothesized to do so by influencing cellulose biosynthesis (Harpaz-Saad et al., 2011, 2012).Arabidopsis (Arabidopsis thaliana) seed coat mucilage is a powerful model for studying cell wall biosynthesis and polysaccharide interactions (Arsovski et al., 2010; Haughn and Western, 2012). Seed coat epidermal cells sequentially produce two distinct types of secondary cell walls with unique morphologies and properties (Western et al., 2000; Windsor et al., 2000). Between approximately 5 and 9 d approximate time of fertilization (DPA), seed coat epidermal cells synthesize mucilage and deposit it in the apoplast, creating a donut-shaped mucilage pocket that surrounds a central cytoplasmic column (Western et al., 2000, 2004; Haughn and Chaudhury, 2005). From 9 to 13 DPA, the cytoplasmic column is gradually replaced by a cellulose-rich, volcano-shaped secondary cell wall called the columella (Beeckman et al., 2000; Western et al., 2000; Windsor et al., 2000; Stork et al., 2010; Mendu et al., 2011).Seed mucilage is composed primarily of relatively unbranched rhamnogalacturonan I (RG I) with minor amounts of homogalacturonan (HG), cellulose, and hemicelluloses (for review, see Haughn and Western, 2012). When mucilage is hydrated, it expands rapidly from the apoplastic pocket, forming a halo that surrounds the seed. Mucilage separates into two fractions: a loose nonadherent fraction and an inner adherent fraction that can only be released by vigorous shaking, strong bases, or glycosidases (for review, see North et al., 2014). Galactans and arabinans are also present in mucilage, and their regulation by glycosidases is required for correct mucilage hydration (Dean et al., 2007; Macquet et al., 2007b; Arsovski et al., 2009). For example, β-XYLOSIDASE1 encodes a bifunctional β-d-xylosidase/α-l-arabinofuranosidase required for arabinan modification in mucilage, and β-xylosidase1 mutant seeds have a delayed mucilage release phenotype (Arsovski et al., 2009). MUCILAGE MODIFIED2 (MUM2) encodes a β-d-galactosidase, and mum2 seeds fail to release mucilage when hydrated in water (Dean et al., 2007; Macquet et al., 2007b). MUM2 is believed to modify RG I galactan side chains but may also affect the galactan component of other mucilage components (Dean et al., 2007; Macquet et al., 2007b). Galactans are capable of binding to cellulose in vitro and could affect mucilage hydration through pectin-cellulose interactions (Zykwinska et al., 2005, 2007a, 2007b; Dick-Pérez et al., 2011; Wang et al., 2012), although carbohydrate linkage analysis suggests that the galactan side chains are very short.Several studies indicate that seed mucilage extrusion and expansion are also influenced by methylesterification of HG. For example, both SUBTILISIN-LIKE SER PROTEASE1.7 and PECTIN METHYLESTERASE INHIBITOR6 are required for proper methyl esterification of mucilage (Rautengarten et al., 2008; Saez-Aguayo et al., 2013). Mutations in another gene, FLYING SAUCER1 (FLY1; a transmembrane E3 ubiquitin ligase), reduce the degree of pectin methylesterification in mucilage and cause increased mucilage adherence and defective mucilage extrusion (Voiniciuc et al., 2013). fly1 seeds have disc-like structures at the edge of the mucilage halo, which are outer primary cell wall fragments that detach from the columella during extrusion and are difficult to separate from the adherent mucilage (Voiniciuc et al., 2013).Recently, CELLULOSE SYNTHASE5 (CESA5) and SOS5 were proposed to facilitate cellulose-mediated mucilage adherence (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011). A simple hypothesis for the role of CESA5 in mucilage adherence is that it synthesizes cellulose, which interacts with the mucilage pectin to mediate adherence. Loss of CESA5 function results in a reduction of mucilage cellulose biosynthesis and a less adherent mucilage cell wall matrix (Mendu et al., 2011; Sullivan et al., 2011). The role of SOS5 in mucilage adherence is more difficult to explain. SOS5 null mutations cause a loss-of-adherence phenotype similar to cesa5-1 seeds, suggesting that SOS5 may regulate mucilage adherence by influencing CESA5 function (Harpaz-Saad et al., 2011). However, the mechanism through which SOS5 could influence CESA5 and/or cellulose biosynthesis is not clear.To better understand the role of SOS5 in mucilage adherence and its relationship to CESA5, we thoroughly investigated the seed coat epidermal cell phenotypes of the cesa5-1 and sos5-2 single mutants as well as those of the cesa5-1 sos5-2 double mutant. We also investigated how cellulose, SOS5, and pectin interact to mediate mucilage adherence by constructing double mutants with either cesa5-1 or sos5-2 together with either mum2-1 or fly1. Our results suggest that SOS5 mediates mucilage adherence independently of CESA5. Furthermore, compared with CESA5, SOS5 has a greater influence on mucilage pectin structure, suggesting that SOS5 mediates mucilage adherence through pectins, not cellulose.     

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.     

Arabidopsis 3-Ketoacyl-Coenzyme A Synthase9 Is Involved in the Synthesis of Tetracosanoic Acids as Precursors of Cuticular Waxes,Suberins, Sphingolipids,and Phospholipids

Very-long-chain fatty acids (VLCFAs) with chain lengths from 20 to 34 carbons are involved in diverse biological functions such as membrane constituents, a surface barrier, and seed storage compounds. The first step in VLCFA biosynthesis is the condensation of two carbons to an acyl-coenzyme A, which is catalyzed by 3-ketoacyl-coenzyme A synthase (KCS). In this study, amino acid sequence homology and the messenger RNA expression patterns of 21 Arabidopsis (Arabidopsis thaliana) KCSs were compared. The in planta role of the KCS9 gene, showing higher expression in stem epidermal peels than in stems, was further investigated. The KCS9 gene was ubiquitously expressed in various organs and tissues, including roots, leaves, and stems, including epidermis, silique walls, sepals, the upper portion of the styles, and seed coats, but not in developing embryos. The fluorescent signals of the KCS9::enhanced yellow fluorescent protein construct were merged with those of BrFAD2::monomeric red fluorescent protein, which is an endoplasmic reticulum marker in tobacco (Nicotiana benthamiana) epidermal cells. The kcs9 knockout mutants exhibited a significant reduction in C24 VLCFAs but an accumulation of C20 and C22 VLCFAs in the analysis of membrane and surface lipids. The mutant phenotypes were rescued by the expression of KCS9 under the control of the cauliflower mosaic virus 35S promoter. Taken together, these data demonstrate that KCS9 is involved in the elongation of C22 to C24 fatty acids, which are essential precursors for the biosynthesis of cuticular waxes, aliphatic suberins, and membrane lipids, including sphingolipids and phospholipids. Finally, possible roles of unidentified KCSs are discussed by combining genetic study results and gene expression data from multiple Arabidopsis KCSs.Very-long-chain fatty acids (VLCFAs) are fatty acids of 20 or more carbons in length and are essential precursors of functionally diverse lipids, cuticular waxes, aliphatic suberins, phospholipids, sphingolipids, and seed oils in the Brassicaceae. These lipids are involved in various functions, such as acting as protective barriers between plants and the environment, impermeable barriers to water and ions, energy-storage compounds in seeds, structural components of membranes, and lipid signaling, which is involved in the hypersensitive response (Pollard et al., 2008; Kunst and Samuels, 2009; Franke et al., 2012). VLCFAs are synthesized by the microsomal fatty acid elongase complex, which catalyzes the cyclic addition of a C2 moiety obtained from malonyl-CoA to C16 or C18 acyl-CoA. The fatty acid elongation process has been shown to proceed through a series of four reactions: condensation of the C2 carbon moiety to acyl-CoA by 3-ketoacyl coenzyme A synthase (KCS), reduction of KCS by 3-ketoacyl coenzyme A reductase (KCR), dehydration of 3-hydroxyacyl-CoA by 3-hydroxyacyl-CoA dehydratase (PAS2), and reduction of trans-2,3-enoyl-CoA by trans-2-enoyl-CoA reductase (ECR). Except for KCS isoforms with redundancy, disruption of KCR1, ECR/ECERIFERUM10 (CER10), or PAS2 exhibited severe morphological abnormalities and embryo lethality, suggesting that VLCFA homeostasis is essential for plant developmental processes (Zheng et al., 2005; Bach et al., 2008; Beaudoin et al., 2009).Cuticular waxes that cover plant aerial surfaces are known to be involved in limiting nonstomatal water loss and gaseous exchanges (Boyer et al., 1997; Riederer and Schreiber, 2001), repelling lipophilic pathogenic spores and dust (Barthlott and Neinhuis, 1997), and protecting plants from UV light (Reicosky and Hanover, 1978). VLCFAs that are synthesized in the epidermal cells are either directly used or further modified into aldehydes, alkanes, secondary alcohols, ketones, primary alcohols, and wax esters for the synthesis of cuticular waxes. Reverse genetic analysis and Arabidopsis (Arabidopsis thaliana) epidermal peel microarray analysis (Suh et al., 2005) has enabled the research community to identify the functions of many genes involved in cuticular wax biosynthesis (Kunst and Samuels, 2009): CER1 (Bourdenx et al., 2011; Bernard et al., 2012), WAX2/CER3 (Chen et al., 2003; Rowland et al., 2007; Bernard et al., 2012), and MAH1(Greer et al., 2007; Wen and Jetter, 2009) have been shown to be involved in the decarbonylation pathway to form aldehydes, alkanes, secondary alcohols, and ketones, and acyl-coenzyme A reductase (FAR; Aarts et al., 1997; Rowland et al., 2006) and WSD1 (Li et al., 2008) have been shown to be involved in the decarboxylation pathway for the synthesis of primary alcohols and wax esters. The export of wax precursors to the extracellular space is mediated by a heterodimer of the ATP-binding cassette transporters in the plasma membrane (Pighin et al., 2004; Bird et al., 2007; McFarlane et al., 2010). In addition, glycosylphosphatidylinositol-anchored LTP (LTPG1) and LTPG2 contribute either directly or indirectly to the export of cuticular wax (DeBono et al., 2009; Lee et al., 2009; Kim et al., 2012).VLCFAs that are synthesized in the endodermis of primary roots, seed coats, and the chalaza-micropyle region of seeds are used as precursors for the synthesis of aliphatic suberins. The suberin layer is known to function as a barrier against uncontrolled water, gas, and ion loss and provides protection from environmental stresses and pathogens (Pollard et al., 2008; Franke et al., 2012). For aliphatic suberin biosynthesis, the ω-carbon of the VLCFAs is oxidized by the fatty acyl ω-hydroxylase (Xiao et al., 2004; Li et al., 2007; Höfer et al., 2008; Molina et al., 2008, 2009; Compagnon et al., 2009; Li-Beisson et al., 2009), and the ω-hydroxy VLCFAs are further oxidized into α,ω-dicarboxylic acids by the HOTHEAD-like oxidoreductase (Kurdyukov et al., 2006). α,ω-Dicarboxylic acids are acylated to glycerol-3-P via acyl-CoA:glycerol-3-P acyltransferase (Beisson et al., 2007; Li et al., 2007; Li-Beisson et al., 2009; Yang et al., 2010) or to ferulic acid. In addition, C18, C20, and C22 fatty acids are also reduced by FAR enzymes to primary fatty alcohols, which are a common component in root suberin (Vioque and Kolattukudy, 1997). Finally, the aliphatic suberin precursors are likely to be extensively polymerized and cross linked with the polysaccharides or lignins in the cell wall.In addition, VLCFAs are found in sphingolipids, including glycosyl inositolphosphoceramides, glycosylceramides, and ceramides and phospholipids, such as phosphatidylethanolamine (PE) and phosphatidyl-Ser (PS), which are present in the extraplastidial membrane (Pata et al., 2010; Yamaoka et al., 2011). For sphingolipid biosynthesis, VLCFA-CoAs and Ser are condensed to form 3-keto-sphinganine, which is subsequently reduced to produce sphinganine, a long chain base (LCB). LCBs are known to be further modified by 4-hydroxylation, 4-desaturation, and 8-desaturation (Lynch and Dunn, 2004; Chen et al., 2006, 2012; Pata et al., 2010). The additional VLCFAs are linked with 4-hydroxy LCBs via an amino group to form ceramides (Chen et al., 2008). The presence of VLCFA in sphingolipids may contribute to an increase of their hydrophobicity, membrane leaflet interdigitation, and the transition from a fluid to a gel phase, which is required for microdomain formation. In plants, PS is synthesized from CDP-diacylglycerol and Ser by PS synthase or through an exchange reaction between a phospholipid head group and Ser by a calcium-dependent base-exchange-type PS synthase (Vincent et al., 1999; Yamaoka et al., 2011). PE biosynthesis proceeds through decarboxylation via PS decarboxylase (Nerlich et al., 2007), the phosphoethanolamine transfer from CDP-ethanolamine to diacylglycerol (Kennedy pathway), and the exchange of the head group of PE with Ser via a base-exchange enzyme (Marshall and Kates, 1973). In particular, PS containing a relatively large amount of VLCFAs is enriched in endoplasmic reticulum (ER)-derived vesicles that may function in stabilizing small (70- to 80-nm-diameter) vesicles (Vincent et al., 2001).During the fatty acid elongation process, the first committed step is the condensation of C₂ units to acyl-CoA by KCS. Arabidopsis harbors a large family containing 21 KCS members (Joubès et al., 2008). Characterization of Arabidopsis KCS mutants with defects in VLCFA synthesis revealed in planta roles and substrate specificities (based on differences in carbon chain length and degree of unsaturation) of KCSs. For example, FAE1, a seed-specific condensing enzyme, was shown to catalyze C20 and C22 VLCFA biosynthesis for seed storage lipids (James et al., 1995). KCS6/CER6/CUT1 and KCS5/CER60 are involved in the elongation of fatty acyl-CoAs longer than C28 VLCFA for cuticular waxes in epidermis and pollen coat lipids (Millar et al., 1999; Fiebig et al., 2000; Hooker et al., 2002). KCS20 and KCS2/DAISY are functionally redundant in the two-carbon elongation to C22 VLCFA, which is required for cuticular wax and root suberin biosynthesis (Franke et al., 2009; Lee et al., 2009). When KCS1 and KCS9 were expressed in yeast (Saccharomyces cerevisiae), KCS1 showed broad substrate specificity for saturated and monounsaturated C16 to C24 acyl-CoAs and KCS9 utilized the C16 to C22 acyl-CoAs (Trenkamp et al., 2004; Blacklock and Jaworski, 2006; Paul et al., 2006). Recently, CER2 encoding putative BAHD acyltransferase was reported to be a fatty acid elongase that was involved in the elongation of C28 fatty acids for the synthesis of wax precursors (Haslam et al., 2012).In this study, the expression patterns and subcellular localization of KCS9 were examined, and an Arabidopsis kcs9 mutant was isolated to investigate the roles of KCS9 in planta. Diverse classes of lipids, including cuticular waxes, aliphatic suberins, and sphingolipids, as well as fatty acids in various organs were analyzed from the wild type, the kcs9 mutant, and complementation lines. The combined results of this study revealed that KCS9 is involved in the elongation of C22 to C24 fatty acids, which are essential precursors for the biosynthesis of cuticular waxes, aliphatic suberins, and membrane lipids, including sphingolipids. To the best of our knowledge, this is the first study where a KCS9 isoform involved in sphingolipid biosynthesis was identified.