Rice Aldehyde Dehydrogenase7 Is Needed for Seed Maturation and Viability

Aldehyde dehydrogenases (ALDHs) catalyze the irreversible oxidation of a wide range of reactive aldehydes to their corresponding carboxylic acids. Although the proteins have been studied from various organisms and at different growth stages, their roles in seed development have not been well elucidated. We obtained T-DNA insertional mutants in OsALDH7, which is remarkably inducible by oxidative and abiotic stresses. Interestingly, endosperms from the osaldh7 null mutants accumulated brown pigments during desiccation and storage. Extracts from the mutant seeds showed a maximum absorbance peak at 360 nm, the wavelength that melanoidin absorbs. Under UV light, those extracts also exhibited much stronger fluorescence than the wild type, suggesting that the pigments are melanoidin. These pigments started to accumulate in the late seed developmental stage, the time when OsALDH7 expression began to increase significantly. Purified OsALDH7 protein showed enzyme activities to malondialdehyde, acetaldehyde, and glyceraldehyde. These results suggest that OsALDH7 is involved in removing various aldehydes formed by oxidative stress during seed desiccation. The mutant seeds were more sensitive to our accelerated aging treatment and accumulated more malondialdehyde than the wild type. These data imply that OsALDH7 plays an important role in maintaining seed viability by detoxifying the aldehydes generated by lipid peroxidation.The major regulatory factors that control seed aging are oxidative stress, lipid peroxidation, and respiration (Sun and Leopold, 1995; Bailly et al., 1996; Akimoto et al., 2004). Lipid peroxidation and respiration result in the formation of reactive aldehydes such as malondialdehyde (MDA) and acetaldehyde, which tend to react with proteins and amino acids (Mueller, 1998; Almeras et al., 2003; Weber et al., 2004). Those reactions cause aging and seed damage (Zhang et al., 1995, 1997). Until recently, a physiological approach has been taken in research on seed aging, and molecular and genetic studies have been seldom reported (Clerkx et al., 2004a).Rice (Oryza sativa) is an important food crop, especially in Asia. Although stress tolerance has been extensively evaluated in an effort to develop advanced cultivars, the aging of seeds is an important economic problem that has been rarely examined (Devaiah et al., 2007). Their deterioration by lipid peroxidation leads to undesirable taste, color, and odor (Robertson et al., 1973; Nakayama et al., 1981; List et al., 1992). This is caused by various reactive aldehydes containing MDA, which are volatile aromatic compounds. Those aldehydes also bind with proteins and amino acids nonenzymatically, resulting in the accumulation of brown pigments during seed storage (Sun and Leopold, 1995).Aldehydes are intermediates in several fundamental metabolism pathways for carbohydrates, vitamins, steroids, amino acids, and lipids (Yoshida et al., 1998). They are also produced in response to environmental stresses that disturb metabolism, including salinity, dehydration, desiccation, cold, and heat shock (Bartels, 2001). Although indispensable for an organism, excessive amounts threaten seed survival because of their chemically reactive nature and the toxic effect of the molecules (Lindahl, 1992). Therefore, aldehyde levels in cells must be regulated tightly.Aldehyde dehydrogenases (ALDHs) are represented by a protein super-family that can be categorized into 21 families in eukaryotes (Perozich et al., 1999; Sophos et al., 2001; Sophos and Vasiliou, 2003; Fong et al., 2006). Some ALDHs are substrate specific while others have a highly variable substrate specificity (Sophos and Vasiliou, 2003). ALDHs catalyze the irreversible oxidation of a wide range of reactive aldehydes to their corresponding carboxylic acids (Perozich et al., 1999; Kirch et al., 2005). Therefore, they play a pivotal role in detoxifying the aldehydes generated by environmental stresses. Plant enzymes are represented in 11 ALDH families (Kirch et al., 2004). Despite their importance, the physiological functions of ALDHs in plants have rarely been studied (Liu et al., 2001; Liu and Schnable, 2002; Sunkar et al., 2003; Tsuji et al., 2003).Family 7 ALDHs (antiquitins) are comparatively distinguishable from other ALDH families, because they show low sequence identity (approximately 30%) to ALDHs from other families (Vaciliou et al., 1999). However, the amino acid sequences among Family 7 members are highly homologous (about 60%–70%; Lee et al., 1994; Fong et al., 2006). In contrast to animal antiquitins that do not exhibit any inducible response to stresses (Lee et al., 1994; Fong et al., 2006), antiquitin in garden pea (Pisum sativum) is inducible by dehydration (Guerrero et al., 1990).Overexpression of Arabidopsis (Arabidopsis thaliana) and soybean (Glycine max) ALDH7 confers tolerance to osmotic and oxidative stresses in transgenic plants (Kotchoni et al., 2006; Rodrigues et al., 2006). Moreover, their MDA and hydrogen peroxide contents are decreased. This suggests that ALDH7s function not only as aldehyde-detoxifying enzymes but also as efficient scavengers of reactive oxygen species and as lipid peroxidation-inhibiting enzymes.In this study, we used null mutants in the rice ALDH7 gene to investigate the functional roles of ALDH7 during seed development and storage.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

CELLULOSE SYNTHASE-LIKE A2, a Glucomannan Synthase,Is Involved in Maintaining Adherent Mucilage Structure in Arabidopsis Seed

Mannans are hemicellulosic polysaccharides that are considered to have both structural and storage functions in the plant cell wall. However, it is not yet known how mannans function in Arabidopsis (Arabidopsis thaliana) seed mucilage. In this study, CELLULOSE SYNTHASE-LIKE A2 (CSLA2; At5g22740) expression was observed in several seed tissues, including the epidermal cells of developing seed coats. Disruption of CSLA2 resulted in thinner adherent mucilage halos, although the total amount of the adherent mucilage did not change compared with the wild type. This suggested that the adherent mucilage in the mutant was more compact compared with that of the wild type. In accordance with the role of CSLA2 in glucomannan synthesis, csla2-1 mucilage contained 30% less mannosyl and glucosyl content than did the wild type. No appreciable changes in the composition, structure, or macromolecular properties were observed for nonmannan polysaccharides in mutant mucilage. Biochemical analysis revealed that cellulose crystallinity was substantially reduced in csla2-1 mucilage; this was supported by the removal of most mucilage cellulose through treatment of csla2-1 seeds with endo-β-glucanase. Mutation in CSLA2 also resulted in altered spatial distribution of cellulose and an absence of birefringent cellulose microfibrils within the adherent mucilage. As with the observed changes in crystalline cellulose, the spatial distribution of pectin was also modified in csla2-1 mucilage. Taken together, our results demonstrate that glucomannans synthesized by CSLA2 are involved in modulating the structure of adherent mucilage, potentially through altering cellulose organization and crystallization.Mannan polysaccharides are a complex set of hemicellulosic cell wall polymers that are considered to have both structural and storage functions. Based on the particular chemical composition of the backbone and the side chains, mannan polysaccharides are classified into four types: pure mannan, glucomannan, galactomannan, and galactoglucomannan (Moreira and Filho, 2008; Wang et al., 2012; Pauly et al., 2013). Each of these polysaccharides is composed of a β-1,4-linked backbone containing Man or a combination of Glc and Man residues. In addition, the mannan backbone can be substituted with side chains of α-1,6-linked Gal residues. Mannan polysaccharides have been proposed to cross link with cellulose and other hemicelluloses via hydrogen bonds (Fry, 1986; Iiyama et al., 1994; Obel et al., 2007; Scheller and Ulvskov, 2010). Furthermore, it has been reported that heteromannans with different levels of substitution can interact with cellulose in diverse ways (Whitney et al., 1998). Together, these observations indicate the complexity of mannan polysaccharides in the context of cell wall architecture.CELLULOSE SYNTHASE-LIKE A (CSLA) enzymes have been shown to have mannan synthase activity in vitro. These enzymes polymerize the β-1,4-linked backbone of mannans or glucomannans, depending on the substrates (GDP-Man and/or GDP-Glc) provided (Richmond and Somerville, 2000; Liepman et al., 2005, 2007; Pauly et al., 2013). In Arabidopsis (Arabidopsis thaliana), nine CSLA genes have been identified; different CSLAs are responsible for the synthesis of different mannan types (Liepman et al., 2005, 2007). CSLA7 has mannan synthase activity in vitro (Liepman et al., 2005) and has been shown to synthesize stem glucomannan in vivo (Goubet et al., 2009). Disrupting the CSLA7 gene results in defective pollen growth and embryo lethality phenotypes in Arabidopsis, indicating structural or signaling functions of mannan polysaccharides during plant embryo development (Goubet et al., 2003). A mutation in CSLA9 results in the inhibition of Agrobacterium tumefaciens-mediated root transformation in the rat4 mutant (Zhu et al., 2003). CSLA2, CSLA3, and CSLA9 are proposed to play nonredundant roles in the biosynthesis of stem glucomannans, although mutations in CSLA2, CSLA3, or CSLA9 have no effect on stem development or strength (Goubet et al., 2009). All of the Arabidopsis CSLA proteins have been shown to be involved in the biosynthesis of mannan polysaccharides in the plant cell wall (Liepman et al., 2005, 2007), although the precise physiological functions of only CSLA7 and CSLA9 have been conclusively demonstrated.In Arabidopsis, when mature dry seeds are hydrated, gel-like mucilage is extruded to envelop the entire seed. Ruthenium red staining of Arabidopsis seeds reveals two different mucilage layers, termed the nonadherent and the adherent mucilage layers (Western et al., 2000; Macquet et al., 2007a). The outer, nonadherent mucilage is loosely attached and can be easily extracted by shaking seeds in water. Compositional and linkage analyses suggest that this layer is almost exclusively composed of unbranched rhamnogalacturonan I (RG-I) (>80% to 90%), with small amounts of branched RG-I, arabinoxylan, and high methylesterified homogalacturonan (HG). By contrast, the inner, adherent mucilage layer is tightly attached to the seed and can only be removed by strong acid or base treatment, or by enzymatic digestion (Macquet et al., 2007a; Huang et al., 2011; Walker et al., 2011). As with the nonadherent layer, adherent mucilage is also mainly composed of unbranched RG-I, but with small numbers of arabinan and galactan ramifications (Penfield et al., 2001; Willats et al., 2001; Dean et al., 2007; Macquet et al., 2007a, 2007b; Arsovski et al., 2009; Haughn and Western, 2012). There are also minor amounts of pectic HG in the adherent mucilage, with high methylesterified HG in the external domain compared with the internal domain of the adherent layer (Willats et al., 2001; Macquet et al., 2007a; Rautengarten et al., 2008; Sullivan et al., 2011; Saez-Aguayo et al., 2013). In addition, the adherent mucilage contains cellulose (Blake et al., 2006; Macquet et al., 2007a), which is entangled with RG-I and is thought to anchor the pectin-rich mucilage onto seeds (Macquet et al., 2007a; Harpaz-Saad et al., 2011, 2012; Mendu et al., 2011; Sullivan et al., 2011). As such, Arabidopsis seed mucilage is considered to be a useful model for investigating the biosynthesis of cell wall polysaccharides and how this process is regulated in vivo (Haughn and Western, 2012).Screening for altered seed coat mucilage has led to the identification of several genes encoding enzymes that are involved in the biosynthesis or modification of mucilage components. RHAMNOSE SYNTHASE2/MUCILAGE-MODIFIED4 (MUM4) is responsible for the synthesis of UDP-l-Rha (Usadel et al., 2004; Western et al., 2004; Oka et al., 2007). The putative GALACTURONSYLTRANSFERASE11 can potentially synthesize mucilage RG-I or HG pectin from UDP-d-GalUA (Caffall et al., 2009). GALACTURONSYLTRANSFERASE-LIKE5 appears to function in the regulation of the final size of the mucilage RG-I (Kong et al., 2011, 2013). Mutant seeds defective in these genes display reduced thickness of the extruded mucilage layer compared with wild-type Arabidopsis seeds.RG-I deposited in the apoplast of seed coat epidermal cells appears to be synthesized in a branched form that is subsequently modified by enzymes in the apoplast. MUM2 encodes a β-galactosidase that removes Gal residues from RG-I side chains (Dean et al., 2007; Macquet et al., 2007b). β-XYLOSIDASE1 encodes an α-l-arabinfuranosidase that removes Ara residues from RG-I side chains (Arsovski et al., 2009). Disruptions of these genes lead to defective hydration properties and affect the extrusion of mucilage. Furthermore, correct methylesterification of mucilage HG is also required for mucilage extrusion. HG is secreted into the wall in a high methylesterified form that can then be enzymatically demethylesterified by pectin methylesterases (PMEs; Bosch and Hepler, 2005). PECTIN METHYLESTERASE INHIBITOR6 (PMEI6) inhibits PME activities (Saez-Aguayo et al., 2013). The subtilisin-like Ser protease (SBT1.7) can activate other PME inhibitors, but not PMEI6 (Rautengarten et al., 2008; Saez-Aguayo et al., 2013). Disruption of either PMEI6 or SBT1.7 results in the delay of mucilage release.Although cellulose is present at low levels in adherent mucilage, it plays an important adhesive role for the attachment of mucilage pectin to the seed coat epidermal cells. The orientation and amount of pectin associated with the cellulose network is largely determined by cellulose conformation properties (Macquet et al., 2007a; Haughn and Western, 2012). Previous studies have demonstrated that CELLULOSE SYNTHASE A5 (CESA5) is required for the production of seed mucilage cellulose and the adherent mucilage in the cesa5 mutant can be easily extracted with water (Harpaz-Saad et al., 2011, 2012; Mendu et al., 2011; Sullivan et al., 2011).Despite all of these discoveries, large gaps remain in the current knowledge of the biosynthesis and functions of mucilage polysaccharides in seed coats. In this study, we show that CSLA2 is involved in the biosynthesis of mucilage glucomannan. Furthermore, we show that CSLA2 functions in the maintenance of the normal structure of the adherent mucilage layer through modifying the mucilage cellulose ultrastructure.     

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.     

The K-Segment of Maize DHN1 Mediates Binding to Anionic Phospholipid Vesicles and Concomitant Structural Changes

Dehydrins (DHNs; late embryogenesis abundant D11 family) are a family of intrinsically unstructured plant proteins that accumulate in the late stages of seed development and in vegetative tissues subjected to water deficit, salinity, low temperature, or abscisic acid treatment. We demonstrated previously that maize (Zea mays) DHNs bind preferentially to anionic phospholipid vesicles; this binding is accompanied by an increase in α-helicity of the protein, and adoption of α-helicity can be induced by sodium dodecyl sulfate. All DHNs contain at least one “K-segment,” a lysine-rich 15-amino acid consensus sequence. The K-segment is predicted to form a class A2 amphipathic α-helix, a structural element known to interact with membranes and proteins. Here, three K-segment deletion proteins of maize DHN1 were produced. Lipid vesicle-binding assays revealed that the K-segment is required for binding to anionic phospholipid vesicles, and adoption of α-helicity of the K-segment accounts for most of the conformational change of DHNs upon binding to anionic phospholipid vesicles or sodium dodecyl sulfate. The adoption of structure may help stabilize cellular components, including membranes, under stress conditions.When plants encounter environmental stresses such as drought or low temperature, various responses take place to adapt to these conditions. Typical responses include increased expression of chaperones, signal transduction pathway and late embryogenesis abundant (LEA) proteins, osmotic adjustment, and induction of degradation and repair systems (Ingram and Bartels, 1996).Dehydrins (DHNs; LEA D11 family) are a subfamily of group 2 LEA proteins that accumulate to high levels during late stages of seed development and in vegetative tissues subjected to water deficit, salinity, low temperature, or abscisic acid (ABA) treatment (Svensson et al., 2002). Some DHNs are expressed constitutively during normal growth (Nylander et al., 2001; Rorat et al., 2004, 2006; Rodriguez et al., 2005). DHNs exist in a wide range of photosynthetic organisms, including angiosperms, gymnosperms, algae, and mosses (Svensson et al., 2002). DHNs are encoded by a dispersed multigene family and are differentially regulated, at least in higher plants. For example, 13 Dhn genes have been identified in barley (Hordeum vulgare), dispersed over seven genetic map locations (Choi et al., 1999; Svensson et al., 2002) and regulated variably by drought, low temperature, and embryo development (Tommasini et al., 2008). DHNs are localized in various subcellular compartments, including cytosol (Roberts et al., 1993), nucleus (Houde et al., 1995), chloroplast (Artus et al., 1996), vacuole (Heyen et al., 2002), and proximal to the plasma membrane and protein bodies (Asghar et al., 1994; Egerton-Warburton et al., 1997; Puhakainen et al., 2004). Elevated expression of Dhn genes generally has been correlated with the acquisition of tolerance to abiotic stresses such as drought (Whitsitt et al., 1997), salt (Godoy et al., 1994; Jayaprakash et al., 1998), chilling (Ismail et al., 1999a), or freezing (Houde et al., 1995; Danyluk et al., 1998; Fowler et al., 2001). The differences in expression and tissue location suggest that individual members of the Dhn multigene family have somewhat distinct biological functions (Close, 1997; Zhu et al., 2000; Nylander et al., 2001). Many studies have observed a positive correlation between the accumulation of DHNs and tolerance to abiotic stresses (Svensson et al., 2002). However, overexpression of a single DHN protein has not, in general, been sufficient to confer stress tolerance (Puhakainen et al., 2004).DHNs are subclassified by sequence motifs referred to as the K-segment (Lys-rich consensus sequence), the Y-segment (N-terminal conserved sequence), the S-segment (a tract of Ser residues), and the φ-segment (Close, 1996). Because of high hydrophilicity, high content of Gly (>20%), and the lack of a defined three-dimensional structure in the pure form (Lisse et al., 1996), DHNs have been categorized as “intrinsically disordered/unstructured proteins” or “hydrophilins” (Wright and Dyson, 1999; Garay-Arroyo et al., 2000; Tompa, 2005; Kovacs et al., 2008). On the basis of compositional and biophysical properties and their link to abiotic stresses, several functions of DHNs have been proposed, including ion sequestration (Roberts et al., 1993), water retention (McCubbin et al., 1985), and stabilization of membranes or proteins (Close, 1996, 1997). Observations from in vitro experiments include DHN binding to lipid vesicles (Koag et al., 2003; Kovacs et al., 2008) or metals (Svensson et al., 2000; Heyen et al., 2002; Kruger et al., 2002; Alsheikh et al., 2003; Hara et al., 2005), protection of membrane lipid against peroxidation (Hara et al., 2003), retention of hydration or ion sequestration (Bokor et al., 2005; Tompa et al., 2006), and chaperone activity against the heat-induced inactivation and aggregation of various proteins (Kovacs et al., 2008).Intrinsically disordered/unstructured proteins that lack a well-defined three-dimensional structure have recently been recognized to be prevalent in prokaryotes and eukaryotes (Oldfield et al., 2005). They fulfill important functions in signal transduction, gene expression, and binding to targets such as protein, RNA, ions, and membranes (Wright and Dyson, 1999; Tompa, 2002; Dyson and Wright, 2005). The disorder confers structural flexibility and malleability to adapt to changes in the protein environment, including water potential, pH, ionic strength, and temperature, and to undergo structural transition when complexed with ligands such as other proteins, DNA, RNA, or membranes (Prestrelski et al., 1993; Uversky, 2002). Structural changes from disorder to ordered functional structure also can be induced by the folding of a partner protein (Wright and Dyson, 1999; Tompa, 2002; Mouillon et al., 2008).The idea that DHNs interact with membranes is consistent with many immunolocalization studies, which have shown that DHNs accumulate near the plasma membrane or membrane-rich areas surrounding lipid and protein bodies (Asghar et al., 1994; Egerton-Warburton et al., 1997; Danyluk et al., 1998; Puhakainen et al., 2004). The K-segment is predicted to form a class A2 amphipathic α-helix, in which hydrophilic and hydrophobic residues are arranged on opposite faces (Close, 1996). The amphipathic α-helix is a structural element known to interact with membranes and proteins (Epand et al., 1995). Also, in the presence of helical inducers such as SDS and trifluoroethanol (Dalal and Pio, 2006), DHNs take on α-helicity (Lisse et al., 1996; Ismail et al., 1999b). We previously examined the binding of DHN1 to liposomes and found that DHNs bind preferentially to anionic phospholipids and that this binding is accompanied by an increase in α-helicity of the protein (Koag et al., 2003). Similarly, a mitochondrial LEA protein, one of the group III LEA proteins, recently has been shown to interact with and protect membranes subjected to desiccation, coupled with the adoption of amphipathic α-helices (Tolleter et al., 2007).Here, we explore the basis of DHN-vesicle interaction using K-segment deletion proteins. This study reveals that the K-segment is necessary and sufficient for binding to anionic phospholipid vesicles and that the adoption of α-helicity of DHN proteins can be attributed mainly to the K-segment.     

RNA Silencing of Exocyst Genes in the Stigma Impairs the Acceptance of Compatible Pollen in Arabidopsis

Initial pollen-pistil interactions in the Brassicaceae are regulated by rapid communication between pollen grains and stigmatic papillae and are fundamentally important, as they are the first step toward successful fertilization. The goal of this study was to examine the requirement of exocyst subunits, which function in docking secretory vesicles to sites of polarized secretion, in the context of pollen-pistil interactions. One of the exocyst subunit genes, EXO70A1, was previously identified as an essential factor in the stigma for the acceptance of compatible pollen in Arabidopsis (Arabidopsis thaliana) and Brassica napus. We hypothesized that EXO70A1, along with other exocyst subunits, functions in the Brassicaceae dry stigma to deliver cargo-bearing secretory vesicles to the stigmatic papillar plasma membrane, under the pollen attachment site, for pollen hydration and pollen tube entry. Here, we investigated the functions of exocyst complex genes encoding the remaining seven subunits, SECRETORY3 (SEC3), SEC5, SEC6, SEC8, SEC10, SEC15, and EXO84, in Arabidopsis stigmas following compatible pollinations. Stigma-specific RNA-silencing constructs were used to suppress the expression of each exocyst subunit individually. The early postpollination stages of pollen grain adhesion, pollen hydration, pollen tube penetration, seed set, and overall fertility were analyzed in the transgenic lines to evaluate the requirement of each exocyst subunit. Our findings provide comprehensive evidence that all eight exocyst subunits are necessary in the stigma for the acceptance of compatible pollen. Thus, this work implicates a fully functional exocyst complex as a component of the compatible pollen response pathway to promote pollen acceptance.In flowering plants, sexual reproduction occurs as a result of constant communication between the male gametophyte and the female reproductive organ, from the initial acceptance of compatible pollen to final step of successful fertilization (for review, see Beale and Johnson, 2013; Dresselhaus and Franklin-Tong, 2013; Higashiyama and Takeuchi, 2015). In the Brassicaceae, the stigmas that present a receptive surface for pollen are categorized as dry and covered with unicellular papillae (Heslop-Harrison and Shivanna, 1977). Communication is initiated rapidly following contact of a pollen grain with a stigmatic papilla, as the role of the papillae is to regulate the early cellular responses leading to compatible pollen germination. The basal compatible pollen recognition response also presents a barrier to foreign pollen or is inhibited with self-incompatible pollen (for review, see Dickinson, 1995; Hiscock and Allen, 2008; Chapman and Goring, 2010; Indriolo et al., 2014b).The initial adhesive interaction between the pollen grain and the papilla cell in the Brassicaceae is mediated by the exine of the pollen grain and the surface of the stigmatic papilla (Preuss et al., 1993; Zinkl et al., 1999). A stronger connection results between the adhered pollen grain and the stigmatic papilla with the formation of a lipid-protein interface (foot) derived from the pollen coat and the stigmatic papillar surface (Mattson et al., 1974; Stead et al., 1980; Gaude and Dumas, 1986; Elleman and Dickinson, 1990; Elleman et al., 1992; Preuss et al., 1993; Mayfield et al., 2001). It is at this point that a Brassicaceae-specific recognition of compatible pollen is proposed to occur (Hülskamp et al., 1995; Pruitt, 1999), though the nature of this recognition system is not clearly defined. Two stigma-specific Brassica oleracea glycoproteins, the S-Locus Glycoprotein and S-Locus Related1 (SLR1) protein, play a role in compatible pollen adhesion (Luu et al., 1997, 1999), potentially through interactions with the pollen coat proteins, PCP-A1 and SLR1-BP, respectively (Doughty et al., 1998; Takayama et al., 2000). The simultaneous recognition of self-incompatible pollen would also take place at this stage (for review, see Dresselhaus and Franklin-Tong, 2013; Indriolo et al., 2014b; Sawada et al., 2014). Thus, this interface not only provides a strengthened bond between the pollen grain and stigmatic papilla, but likely facilitates the interaction of signaling proteins from both partners to promote specific cellular responses in the stigmatic papilla toward the pollen grain.One response regulated by these interactions is the release of water from the stigmatic papilla to the adhered compatible pollen grain to enable the pollen grain to rehydrate, germinate, and produce a pollen tube (Zuberi and Dickinson, 1985; Preuss et al., 1993). Upon hydration, the pollen tube emerges at the site of pollen-papilla contact and penetrates the stigma surface between the plasma membrane and the overlaying cell wall (Elleman et al., 1992; Kandasamy et al., 1994). Pollen tube entry into the stigmatic surface represents a second barrier, selecting compatible pollen tubes. Subsequently, the compatible pollen tubes traverse down to the base of the stigma, enter the transmitting tract, and grow intracellularly toward ovules for fertilization. Pollen-pistil interactions at these later stages are also highly regulated (for review, see Beale and Johnson, 2013; Dresselhaus and Franklin-Tong, 2013; Higashiyama and Takeuchi, 2015).EXO70A1, a subunit of the exocyst, was identified as a factor involved in early pollen-stigma interactions, where it is required in the stigma for the acceptance of compatible pollen and inhibited by the self-incompatibility response (Samuel et al., 2009). Stigmas from the Arabidopsis (Arabidopsis thaliana) exo70A1 mutant display constitutive rejection of wild-type-compatible pollen (Samuel et al., 2009; Safavian et al., 2014). This stigmatic defect was rescued by the stigma-specific expression of an Red Fluorescent Protein (RFP):EXO70A1 transgene (Samuel et al., 2009) or partially rescued by providing a high relative humidity environment (Safavian et al., 2014). In addition, the stigma-specific expression of an EXO70A1 RNA interference construct in Brassica napus ‘Westar’ resulted in impaired compatible pollen acceptance and a corresponding reduction in seed production compared with compatible pollinations with wild-type B. napus ‘Westar’ pistils (Samuel et al., 2009). From these studies, EXO70A1 was found to be a critical component in stigmatic papillae to promote compatible pollen hydration and pollen tube entry through the stigma surface. One of the functions of the exocyst is to mediate polar secretion (for review, see Heider and Munson, 2012; Zárský et al., 2013; Synek et al., 2014). Consistent with this, previous studies have observed vesicle-like structures in proximity to the stigmatic papillar plasma membrane in response to compatible pollen in both Brassica spp. and Arabidopsis species (Elleman and Dickinson, 1990, 1996; Dickinson, 1995; Safavian and Goring, 2013; Indriolo et al., 2014a). The secretory activity is predicted to promote pollen hydration and pollen tube entry. As well, consistent with the proposed inhibition of EXO70A1 by the self-incompatibility pathway (Samuel et al., 2009), a complete absence or a significant reduction of vesicle-like structures at the stigmatic papillar plasma membrane was observed in the exo70A1 mutant and with self-incompatible pollen (Safavian and Goring, 2013; Indriolo et al., 2014a).The exocyst is a well-defined complex in yeast (Saccharomyces cerevisiae) and animal systems, consisting of eight subunits, SEC3, SEC5, SEC6, SEC8, SEC10, SEC15, EXO70, and EXO84 (TerBush et al., 1996; Guo et al., 1999). Exocyst subunit mutants were first identified in yeast as secretory mutants displaying a cytosolic accumulation of secretory vesicles (Novick et al., 1980). Subsequent work defined roles for the exocyst in vesicle docking at target membranes in processes such as regulated secretion, polarized exocytosis, and cytokinesis to facilitate membrane fusion by Soluble NSF Attachment protein Receptor (SNARE) complexes (for review, see Heider and Munson, 2012; Liu and Guo, 2012). In plants, genes encoding all eight exocyst subunits have been identified, and many of these genes exist as multiple copies. For example, the Arabidopsis genome contains single copy genes for SEC6 and SEC8, two copies each for SECRETORY3 (SEC3), SEC5, SEC10, and SEC15, three EXO84 genes, and 23 EXO70 genes (Chong et al., 2010; Cvrčková et al., 2012; Vukašinović et al., 2014). Ultrastructural studies using electron tomography uncovered the existence of a structure resembling the exocyst in Arabidopsis (Otegui and Staehelin, 2004; Seguí-Simarro et al., 2004). Localization studies of specific Arabidopsis exocyst subunits also supported conserved roles in polarized exocytosis and cytokinesis in plants. Localization studies have shown EXO70, SEC6, and SEC8 at the growing tip of pollen tubes (Hála et al., 2008), EXO70A1 at the stigmatic papillar plasma membrane (Samuel et al., 2009), SEC3a, SEC6, SEC8, SEC15b, EXO70A1, and EXO84b at the root epidermal cell plasma membrane and developing cell plate (Fendrych et al., 2010, 2013; Wu et al., 2013; Zhang et al., 2013; Rybak et al., 2014), and SEC3a at the plasma membrane in the embryo and root hair (Zhang et al., 2013). Similar to the yeast exocyst mutants, vesicle accumulation has also been observed in the exo70A1 and exo84b mutants (Fendrych et al., 2010; Safavian and Goring, 2013). Taken together, these findings strongly support that plant exocyst subunits function in vivo in vesicle docking at sites of polarized secretion and cytokinesis (for review, see Zárský et al., 2013). In support of this, a recent study investigating Transport Protein Particle (TRAPP)II and exocyst complexes during cytokinesis in Arabidopsis has identified all eight exocyst components in immunoprecipitated complexes (SEC3a/SEC3b, SEC5a, SEC6, SEC8, SEC10, SEC15b, EXO70A1, EXO70H2, and EXO84b; Rybak et al., 2014).Several plant exocyst subunit genes have been implicated in biological processes that rely on regulated vesicle trafficking, where corresponding mutants have displayed a range of growth defects. At the cellular level, these phenotypes have been associated with decreased cell elongation and polar growth (Cole et al., 2005, 2014; Wen et al., 2005; Synek et al., 2006), defects in cytokinesis and cell plate formation (Fendrych et al., 2010; Wu et al., 2013; Rybak et al., 2014), and disrupted Pin-Formed (PIN) auxin efflux carrier recycling and polar auxin transport (Drdová et al., 2013). Several Arabidopsis subunit mutants display strong growth defects such as the sec3a mutant with an embryo-lethal phenotype (Zhang et al., 2013), sec6, sec8, and exo84b mutants with severely dwarfed phenotypes and defects in root growth (Fendrych et al., 2010; Wu et al., 2013; Cole et al., 2014), and exo70A1 with a milder dwarf phenotype (Synek et al., 2006). The Arabidopsis exo70A1 mutant has also been reported to have defects in root hair elongation, hypocotyl elongation, compatible pollen acceptance, seed coat deposition, and tracheary element differentiation (Synek et al., 2006; Samuel et al., 2009; Kulich et al., 2010; Li et al., 2013). Essential roles for other exocyst subunits include Arabidopsis SEC5a/SEC5b, SEC6, SEC8, and SEC15a/SEC15b in male gametophyte development and pollen tube growth (Cole et al., 2005; Hála et al., 2008; Wu et al., 2013), SEC8 in seed coat deposition (Kulich et al., 2010), SEC5a, SEC8, EXO70A1, and EXO84b in root meristem size and root cell elongation (Cole et al., 2014), and a maize (Zea mays) SEC3 homolog in root hair elongation (Wen et al., 2005). Finally, the Arabidopsis EXO70B1, EXO70B2, and EXO70H1 subunits have been implicated in plant defense responses (Pecenková et al., 2011; Stegmann et al., 2012; Kulich et al., 2013; Stegmann et al., 2013).Even with these detailed studies on the functions of exocyst subunits in plants, a systematic demonstration of the requirement of all eight exocyst subunits in a specific plant biological process is currently lacking. EXO70A1 was previously identified as an essential factor in the stigma for compatible pollen-pistil interactions in Arabidopsis and B. napus (Samuel et al., 2009), and we hypothesized that this protein functions as part of the exocyst complex to tether post-Golgi secretory vesicles to stigmatic papillar plasma membrane (Safavian and Goring, 2013). To provide support for the proposed biological role of the exocyst in the stigma for compatible pollen acceptance, we investigated the roles of the remaining seven subunits, SEC3, SEC5, SEC6, SEC8, SEC10, SEC15, and EXO84, in Arabidopsis stigmatic papillae. Given that some Arabidopsis exocyst subunits were previously determined to be essential at earlier growth stages, stigma-specific RNA-silencing constructs were used for each exocyst subunit, and the early postpollination stages were analyzed for these transgenic lines. Our collective data demonstrates that all eight exocyst subunits are required in the stigma for the early stages of compatible pollen-pistil interactions.