Specialization of Oleosins in Oil Body Dynamics during Seed Development in Arabidopsis Seeds

Oil bodies (OBs) are seed-specific lipid storage organelles that allow the accumulation of neutral lipids that sustain plantlet development after the onset of germination. OBs are covered with specific proteins embedded in a single layer of phospholipids. Using fluorescent dyes and confocal microscopy, we monitored the dynamics of OBs in living Arabidopsis (Arabidopsis thaliana) embryos at different stages of development. Analyses were carried out with different genotypes: the wild type and three mutants affected in the accumulation of various oleosins (OLE1, OLE2, and OLE4), three major OB proteins. Image acquisition was followed by a detailed statistical analysis of OB size and distribution during seed development in the four dimensions (x, y, z, and t). Our results indicate that OB size increases sharply during seed maturation, in part by OB fusion, and then decreases until the end of the maturation process. In single, double, and triple mutant backgrounds, the size and spatial distribution of OBs are modified, affecting in turn the total lipid content, which suggests that the oleosins studied have specific functions in the dynamics of lipid accumulation.The seed is a complex, specific structure that allows a quiescent plant embryo to cope with unfavorable germinating conditions and also permits dissemination of the species. To achieve these functions, seeds accumulate reserve compounds that will ensure the survival of the embryo and fuel the growth of the plantlet upon germination. Accumulation of lipids occurs in many eukaryotic cells and is a rather common means of storing carbon and energy. Lipid droplets (LDs) can be found in all eukaryotes, such as yeast (Saccharomyces cerevisiae; Leber et al., 1994), mammals (Murphy, 2001; Hodges and Wu, 2010), Caenorhabditis elegans (Zhang et al., 2010; Mak, 2012), Drosophila melanogaster (Beller et al., 2006, 2010), and plants (Hsieh and Huang, 2004), but also in prokaryotes (Wältermann et al., 2005). The basic structure of an LD is a core of neutral lipids covered by a phospholipid monolayer. LDs differ between species by the set of proteins covering their surface, the nature of the lipids stored, and their turnover. Nevertheless, they apparently always ensure the same function in the cell (i.e. energy storage; Murphy, 2012). In Brassicacea species such as Arabidopsis (Arabidopsis thaliana), seed reserves are mainly composed of carbohydrates, proteins, and lipids (Baud et al., 2002). The lipids are primarily stored as triacylglycerols (TAGs) in LDs, more commonly called oil bodies (OBs; Hsieh and Huang, 2004; Chapman et al., 2012; Chapman and Ohlrogge, 2012) of diameter 0.5 to 2 µm (Tzen et al., 1993).The protein composition of seed OBs has been determined for several plant species, including Brassica napus (Katavic et al., 2006; Jolivet et al., 2009) and Arabidopsis (Jolivet et al., 2004; D’Andréa et al., 2007; Vermachova et al., 2011). In Arabidopsis, 10 proteins have been identified, and seed-specific oleosins represent up to 79% of the OB proteins (Jolivet et al., 2004; D’Andréa et al., 2007; Vermachova et al., 2011). Oleosins are rather small proteins of 18.5 to 21.2 kD with a specific and highly conserved central hydrophobic domain of 72 amino acid residues flanked by hydrophilic domains of variable size and amino acid composition (Qu and Huang, 1990; Tzen et al., 1990, 1992; Huang, 1996; Hsieh and Huang, 2004). It is generally agreed that oleosins cover the OB surface, with their central hydrophobic domain inserted in the TAG through the phospholipid layer (Tzen and Huang, 1992). Besides their structural function in OBs, oleosins may serve as docking stations for other proteins at its surface (Wilfling et al., 2013) and may participate in the biosynthesis and mobilization of plant oils (Parthibane et al., 2012a, 2012b). Oleosins are probably involved in OB stability (Leprince et al., 1998; Shimada et al., 2008) and in the regulation of OB repulsion (Heneen et al., 2008), preventing the coalescence of OBs into a single organelle (Schmidt and Herman, 2008). Nevertheless, the precise functions of oleosins in OB biogenesis and dynamics have not yet been established.Global analysis of seed lipids can be performed using gas chromatography (Li et al., 2006), which allows the precise determination of both lipid content and fatty acid composition. Recently, direct organelle mass spectrometry has been used to visualize the lipid composition of cotton (Gossypium hirsutum) seed OBs (Horn et al., 2011). Nevertheless, in both cases, the methods are destructive. To observe lipid accumulation at the subcellular level, well-known nondestructive techniques for lipid visualization have been adapted to seeds. Third harmonic generation microscopy (Débarre et al., 2006) and label-free coherent anti-Stokes Raman scattering microscopy (Paar et al., 2012) allow dyeless observation of LDs but require very specific equipment. Magnetic resonance imaging enables topographic analysis of lipid distribution in cereal grains (Neuberger et al., 2008) and in submillimeter-sized seeds like those of tobacco (Nicotiana tabacum; Fuchs et al., 2013). Nevertheless, the use of fluorescent dyes such as Nile Red (Greenspan and Fowler, 1985), BODIPY (Pagano et al., 1991), or LipidTOX (Invitrogen) associated with confocal microscopy is also a powerful way to monitor LDs in living organisms.Despite knowledge accumulated on this topic (Brasaemle and Wolins, 2012; Chapman et al., 2012), little is known about OB dynamics during seed maturation. In this article, we investigate this question by monitoring the evolution of OBs in living Arabidopsis embryos over time. This analysis showed a marked change in OB size at 9 to 10 d after flowering (DAF). We then examined single, double, and triple mutants of the major oleosins found in developing seeds (OLE1 [At4g25140], OLE2 [At5g40420], and OLE4 [At3g01570]; Jolivet et al., 2004). We analyzed the OB dynamics in these mutant backgrounds as if they would contain only these three proteins. We show that the lack of specific oleosins influences the dynamics and distribution of OBs during seed maturation, which in turn affects lipid accumulation. These results pave the way for analyzing specific functions of oleosins in the synthesis, growth, and evolution of OBs.     

Integration of Experiments across Diverse Environments Identifies the Genetic Determinants of Variation in Sorghum bicolor Seed Element Composition

Seedling establishment and seed nutritional quality require the sequestration of sufficient element nutrients. The identification of genes and alleles that modify element content in the grains of cereals, including sorghum (Sorghum bicolor), is fundamental to developing breeding and selection methods aimed at increasing bioavailable element content and improving crop growth. We have developed a high-throughput work flow for the simultaneous measurement of multiple elements in sorghum seeds. We measured seed element levels in the genotyped Sorghum Association Panel, representing all major cultivated sorghum races from diverse geographic and climatic regions, and mapped alleles contributing to seed element variation across three environments by genome-wide association. We observed significant phenotypic and genetic correlation between several elements across multiple years and diverse environments. The power of combining high-precision measurements with genome-wide association was demonstrated by implementing rank transformation and a multilocus mixed model to map alleles controlling 20 element traits, identifying 255 loci affecting the sorghum seed ionome. Sequence similarity to genes characterized in previous studies identified likely causative genes for the accumulation of zinc, manganese, nickel, calcium, and cadmium in sorghum seeds. In addition to strong candidates for these five elements, we provide a list of candidate loci for several other elements. Our approach enabled the identification of single-nucleotide polymorphisms in strong linkage disequilibrium with causative polymorphisms that can be evaluated in targeted selection strategies for plant breeding and improvement.Sorghum (Sorghum bicolor) is a globally cultivated source of food, feed, and fiber. Contrasting needs for elemental nutrient accumulation limit crop yield and quality for sorghum marketed to different sectors. The seed-bearing reproductive organs, or panicles, in sorghum represent up to 30% of the total dry matter yield (Amaducci et al., 2004). Plant-based diets, in which grains compose the major food source, require the accumulation of bioavailable essential elements in the plant seeds. Currently, iron (Fe) and zinc (Zn) deficiencies negatively affect the health of over two billion people worldwide (World Health Organization, 2002). Increased bioavailable elemental nutrient content in the edible portions of sorghum for human and animal nutrition could ameliorate this nutritional crisis (Graham et al., 1999; World Health Organization, 2002). Additional global health benefits could be achieved by increasing magnesium (Mg), selenium (Se), calcium (Ca), and copper (Cu; White and Broadley, 2005) while reducing the concentration of toxic elements, including arsenic (As) and cadmium (Cd; Ma et al., 2008).Seed element accumulation results from interconnected biological processes, including element uptake by the roots, translocation and remobilization within the plant, and ultimately import, deposition, and assimilation/storage in the seeds. Element availability is further affected by the accumulation of metabolites in seeds (Vreugdenhil et al., 2004). High-throughput ionomic analysis, or concurrent measurement of multiple elements, allows for the quantitative and simultaneous measurement of an organism’s elemental composition, providing a snapshot of the functional state of an organism under different experimental conditions (Salt et al., 2008). Most studies of the plant ionome utilize inductively coupled plasma mass spectroscopy (ICP-MS). Briefly, inductively coupled plasma (ICP) functions to ionize the analyte into atoms, which are then detected by mass spectroscopy. Reference standards are used to identify and quantitate each element of interest in the sample. ICP-MS analysis can be accomplished in as little as 1 min per sample, which allows for high-throughput processing of thousands of samples (Salt et al., 2008). Previous studies have demonstrated that several elements, including Fe, manganese (Mn), Zn, cobalt (Co), and Cd, share mechanisms of accumulation (Yi and Guerinot, 1996; Vert et al., 2002; Connolly et al., 2003). Ionomic signatures derived from multiple elements also have been shown to better predict plant physiological status for some elements than the measure of the element’s concentration, including essential nutrients like Fe (Baxter et al., 2008). Holistically examining the ionome provides significant insights into the networks underlying ion homeostasis beyond single-element studies (Baxter and Dilkes, 2012).There are over 45,000 catalogued lines of sorghum at the U.S. Department of Agriculture Germplasm Resource Information Network. This diverse collection of sorghum germplasm contains genetic variation with undiscovered impact on seed element composition (Das et al., 1997). Mapping quantitative trait loci for seed element concentration has been successful in a number of species, including Arabidopsis (Arabidopsis thaliana; Vreugdenhil et al., 2004; Waters and Grusak, 2008; Buescher et al., 2010), rice (Oryza sativa; Norton et al., 2010; Zhang et al., 2014), wheat (Triticum aestivum; Shi et al., 2008; Peleg et al., 2009), and maize (Zea mays; Simić et al., 2012; Baxter et al., 2013, 2014). Genome-wide association (GWA) mapping is well suited for uncovering the genetic basis for complex traits, including seed element accumulation. One of the key strengths of association mapping is that a priori knowledge is not necessary to identify new loci associated with the trait of interest. Furthermore, a GWA mapping population is composed of lines that have undergone numerous recombination events, allowing for a narrower mapping interval. Previous GWA studies in maize (Tian et al., 2011), rice (Huang et al., 2010), and sorghum (Morris et al., 2013) have been successful in identifying the genetic basis for various agronomic traits. Here, we analyzed the seed ionome from a community-generated association panel to identify potential loci underlying seed element accumulation in sorghum.     

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.     

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.