Suberin-Associated Fatty Alcohols in Arabidopsis: Distributions in Roots and Contributions to Seed Coat Barrier Properties

Suberin is found in a variety of tissues, such as root endoderms and periderms, storage tuber periderms, tree cork layer, and seed coats. It acts as a hydrophobic barrier to control the movement of water, gases, and solutes as well as an antimicrobial barrier. Suberin consists of polymerized phenolics, glycerol, and a variety of fatty acid derivatives, including primary fatty alcohols. We have conducted an in-depth analysis of the distribution of the C18:0 to C22:0 fatty alcohols in Arabidopsis (Arabidopsis thaliana) roots and found that only 20% are part of the root suberin polymer, together representing about 5% of its aliphatic monomer composition, while the remaining 80% are found in the nonpolymeric (soluble) fraction. Down-regulation of Arabidopsis FATTY ACYL REDUCTASE1 (FAR1), FAR4, and FAR5, which collectively produce the fatty alcohols found in suberin, reduced their levels by 70% to 80% in (1) the polymeric and nonpolymeric fractions from roots of tissue culture-grown plants, (2) the suberin-associated root waxes from 7-week-old soil-grown plants, and (3) the seed coat suberin polymer. By contrast, the other main monomers of suberin were not altered, indicating that reduced levels of fatty alcohols did not influence the suberin polymerization process. Nevertheless, the 75% reduction in total fatty alcohol and diol loads in the seed coat resulted in increased permeability to tetrazolium salts and a higher sensitivity to abscisic acid. These results suggest that fatty alcohols and diols play an important role in determining the functional properties of the seed coat suberin barrier.Suberin is a cell wall-linked polymeric barrier that plays a critical role in the survival of plants by protecting them against various biotic and abiotic stresses. It primarily acts as a hydrophobic barrier to control the movement of water, gases, and solutes, but also contributes to the strength of the cell wall (Ranathunge et al., 2011). Suberin is deposited at the inner face of primary cell walls next to the plasma membrane (Kolattukudy, 1980; Franke and Schreiber, 2007). It is typically found as lamellae (alternating dark and light bands when viewed by transmission electron microscopy) in the endodermis, exodermis, and peridermis of roots, as well as in the peridermis of underground storage tubers (Bernards, 2002). Suberin is also found in shoot periderms of trees (i.e. cork layer) and in seed coats (Molina et al., 2006, 2008) and is deposited in response to wounding (Kolattukudy, 2001).Suberin is a polymer consisting of aliphatics (fatty acid derivatives), phenolics, and glycerol. The predominant aliphatic components of suberin are ω-hydroxy fatty acids, α,ω-dicarboxylic acids, very-long-chain fatty acids, and primary fatty alcohols, while the major phenolic components are p-hydroxycinnamic acids, especially ferulic acid (Kolatukudy, 1980; Bernards et al., 1995; Pollard et al., 2008; Ranathunge et al., 2011). In the periderm of underground storage organs, suberin is found in association with waxes, which are isolated either by extensive extraction in solvent (Soliday et al., 1979; Serra et al., 2009) or by brief immersion of tubers in chloroform (Espelie et al., 1980). These suberin-associated waxes consist of linear aliphatics (e.g. alkanes, fatty acids, and fatty alcohols), which are similar to cuticular wax components of aerial tissues but generally of shorter chain lengths (Espelie et al., 1980). In waxes extracted from 3-week-old wounded potato (Solanum tuberosum) periderms, alkyl ferulates (i.e. ferulic acid linked by an ester bond to a C16:0–C32:0 fatty alcohol) represent up to 60% of the total wax load (Schreiber et al., 2005). Root waxes are also found in 6- to 7-week-old mature taproots of Arabidopsis (Arabidopsis thaliana) with a fully developed periderm (Li et al., 2007; Kosma et al., 2012). They are enriched in alkyl hydroxycinnamates (AHCs) made of C18:0 to C22:0 fatty alcohols esterified with coumaric, caffeic, or ferulic acids (Kosma et al., 2012). The monomer composition (in terms of major chemical species and chain length) of both suberin and suberin-associated waxes varies considerably between plant species, tissues, and developmental stages. Aliphatic suberin and suberin-associated waxes are considered the major contributors to the diffusion resistance of suberized cell walls to radial transport of water and solutes (Soliday et al., 1979; Espelie et al., 1980; Zimmermann et al., 2000; Ranathunge and Schreiber, 2011). The organization of suberin components in the lamellated structure as well as how waxes may be associated with the polymer is a matter of debate (Graça and Santos, 2007).Primary fatty alcohols are long-chain hydrocarbons containing a single hydroxyl group at the terminal position. They are ubiquitously detected as components of the suberin polymer, representing 1% to 10% of the total monomer mass recovered after transesterification (Holloway, 1983; Bernards, 2002; Pollard et al., 2008). Primary fatty alcohols are also typical components of suberin-associated waxes, where they can be found either in free form or linked by an ester bond with a hydroxycinnamic acid (i.e. as AHCs; Soliday et al., 1979; Espelie et al., 1980; Bernards and Lewis 1992; Li et al., 2007; Kosma et al., 2012). In mechanically isolated endodermis of soybean (Glycine max) roots, fatty alcohols represent about 1.5% and 0.2% of the total aliphatics found in suberin-associated waxes and suberin polymer, respectively (Thomas et al., 2007). In onion (Allium cepa) root exodermis, fatty alcohols (C14:0–C28:0) account for 7% to 12% of the soluble fraction, while the suberin fraction contains only C22:0 fatty alcohol, which makes up 3% of the suberin fraction across all exodermal maturation zones (Meyer et al., 2011). In suberizing potato periderms 7 d post wounding, C16:0 to C28:0 fatty alcohols represent about 10% and 18% of the total aliphatics in the insoluble poly(aliphatic) domain (suberin polymer) and in the soluble (nonpolymeric) fraction, respectively (Yang and Bernards, 2006). A similar study on native periderms from 21-d-stored potato (Serra et al., 2009) reported that fatty alcohols represent about 20% of the total aliphatic components found in the suberin polyester, while unlinked fatty alcohols and alkyl ferulates accounted for about 23% and 44% of the total aliphatics in the soluble waxes.In Arabidopsis, C18:0, C20:0, and C22:0 fatty alcohols account for slightly less than 3% of the polymerized aliphatics in roots of soil-grown plants (Domergue et al., 2010), but as much as 36% [w/w] of the soluble wax load (Li et al., 2007). Arabidopsis fatty acyl reductases FAR1 (At5g22500), FAR4 (At3g44540), and FAR5 (At3g44550) generate, respectively, the C22:0, C20:0, and C18:0 fatty alcohol present in the suberin of root, seed coat, and wounded leaf tissues (Domergue et al., 2010). These three enzymes also generate the C18:0 to C22:0 fatty alcohol components that make up AHCs of root waxes (Kosma et al., 2012). Although one particular chain length of primary alcohol was reduced in each far single mutant line (C18:0-OH, C20:0-OH, and C22:0-OH in far5, far4, and far1, respectively), the total fatty alcohol load of the suberin polymer and its composition were only slightly affected and mutant plants had no obvious developmental or physiological defects (Domergue et al., 2010). In this study, we report on the distribution of primary fatty alcohols in the soluble (nonpolymeric) and insoluble (suberin polymer) fractions from mature roots of Arabidopsis. We report that far double and triple mutants have highly reduced fatty alcohol levels, in a chain length-specific manner, in both fractions as well as in the seed coat suberin polymer. The significant reductions in total fatty alcohol and diol levels in the seed coat of these mutants lead to increased permeability and higher sensitivity to abscisic acid (ABA), bringing to light insights on the roles of fatty alcohols and diols in determining functional properties of suberin.     

CYP86A33-Targeted Gene Silencing in Potato Tuber Alters Suberin Composition,Distorts Suberin Lamellae,and Impairs the Periderm's Water Barrier Function

Suberin is a cell wall lipid polyester found in the cork cells of the periderm offering protection against dehydration and pathogens. Its biosynthesis and assembly, as well as its contribution to the sealing properties of the periderm, are still poorly understood. Here, we report on the isolation of the coding sequence CYP86A33 and the molecular and physiological function of this gene in potato (Solanum tuberosum) tuber periderm. CYP86A33 was down-regulated in potato plants by RNA interference-mediated silencing. Periderm from CYP86A33-silenced plants revealed a 60% decrease in its aliphatic suberin load and greatly reduced levels of C18:1 ω-hydroxyacid (approximately 70%) and α,ω-diacid (approximately 90%) monomers in comparison with wild type. Moreover, the glycerol esterified to suberin was reduced by 60% in the silenced plants. The typical regular ultrastructure of suberin, consisting of dark and light lamellae, disappeared and the thickness of the suberin layer was clearly reduced. In addition, the water permeability of the periderm isolated from CYP86A33-silenced lines was 3.5 times higher than that of the wild type. Thus, our data provide convincing evidence for the involvement of ω-functional fatty acids in establishing suberin structure and function.Periderm, the boundary tissue that replaces the epidermis in the secondary organs of plants, provides efficient protection against dehydration, UV radiation, and pathogens (Esau, 1965). Periderm is regularly found in the bark of woody plants, but herbaceous plants may also form a well-developed periderm in roots, tubers, and the oldest portions of stem. The periderm has been widely studied in potato (Solanum tuberosum) tubers because of the latter''s great agronomic significance (Schmidt and Schönherr, 1982; Vogt et al., 1983; Lulai and Freeman, 2001; Sabba and Lulai, 2002). Shrinkage and flaccidity occur in tubers if the protection afforded by the periderm against water loss is compromised (Lulai et al., 2006). Suberin and waxes embedded into the suberin matrix are considered essential for periderm imperviousness (Franke and Schreiber, 2007). Chemically, suberin is a complex lipid polymer consisting of a fatty acid-derived domain (aliphatic suberin) cross-linked by ester bonds to a polyaromatic lignin-like domain (aromatic suberin; Kolattukudy, 2001; Bernards, 2002; Franke and Schreiber, 2007). Aliphatic suberin has been widely analyzed in potato periderm (Kolattukudy and Agrawal, 1974; Graça and Pereira, 2000; Schreiber et al., 2005). The main monomers released from potato aliphatic suberin are a mixture of ω-hydroxyacids and α,ω-diacids with chain lengths ranging from C16 to C28 (mainly C18), together with glycerol. Small amounts of monofunctional fatty acids, alcohols, and ferulic acid are also released. Waxes are complex mixtures of lipids extractable with chloroform that in potato periderm consist mostly of linear very-long-chain aliphatics up to C32 (Schreiber et al., 2005). Suberin is deposited in the cell wall as a continuous deposit or secondary cell wall that overlays the polysaccharide primary cell wall from the inside (Esau, 1965). These suberin deposits appear under the transmission electron microscope (TEM) as regularly alternating opaque and translucent lamellae (Schmidt and Schönherr, 1982). Although several molecular models for suberin have been proposed (Kolattukudy, 1980; Bernards, 2002; Graça and Santos, 2007), how the suberin and wax components are organized in the lamellated suberin secondary cell wall is a matter of debate (Graça and Santos, 2007). Moreover, to what extent suberin and wax deposition and composition determine sealing properties of periderm still remains unclear (Schreiber et al., 2005). Several studies confirm the importance of waxes for the diffusion barrier (Soliday et al., 1979; Vogt et al., 1983; Schreiber et al., 2005), but the significance of aliphatic suberin has hardly been investigated at all. Interestingly, an Arabidopsis (Arabidopsis thaliana) knockout mutant for the GLYCEROL-3-PHOSPHATE ACYLTRANSFERASE5 gene (GPAT5) with altered suberin showed higher tetrazolium salt permeability in the seed coat (Beisson et al., 2007).ω-Hydroxylation of fatty acids, a reaction carried out in plants by cytochrome P450 monooxygenases, is a crucial step in the biosynthesis of plant biopolymers (Kolattukudy, 1980; Nawrath, 2002). The Arabidopsis mutant lacerata, which shows phenotype defects compatible with a cutin deficiency, is defective in CYP86A8 encoding a fatty acid ω-hydroxylase (Wellesen et al., 2001). The aberrant induction of type three genes1 (att1) mutant, showing an altered cuticle ultrastructure and a higher transpiration rate than wild type, is defective in CYP86A2 and contains reduced amounts of ω-functionalized cutin monomers (Xiao et al., 2004). Moreover, a genome-wide study of cork oak (Quercus suber) bark highlighted a member of the cytochrome P450 of the CYP86A subfamily as a strong candidate gene for aliphatic suberin biosynthesis (Soler et al., 2007); and a role for CYP86A1 in the biosynthesis of suberin has recently been confirmed in the primary root of Arabidopsis knockout mutants (Li et al., 2007; Hofer et al., 2008). However, how the lack of fatty acid ω-hydroxylase activity may affect the structural features and sealing properties of suberin in periderm cell walls has not been documented.To provide experimental evidence of the role of CYP86A genes in periderm fine structure and water transpiration properties, especially quantitative permeability studies so far unexplored in Arabidopsis, we followed a strategy based on the potato (cv Desirée). The potato is a model of choice for such studies because transgenic plants can be produced in reasonable time and sufficient amounts of periderm can easily be obtained from tubers for chemical and physiological studies (Vogt et al., 1983; Schreiber et al., 2005). Based on the CYP86A gene that was highlighted in cork oak periderm as a strong suberin candidate for aliphatic suberin biosynthesis, we isolated the putative ortholog in potato and used a reverse genetic approach to analyze the effects of down-regulation on the chemical and ultrastructural features of suberin and on the physiological properties of the tuber periderm. Our findings indicate that down-regulation of CYP86A33, as this gene was designated, results in a strong decrease of the ω-functionalized monomers in aliphatic suberin, which are necessary for the suberin typical lamellar organization and for the periderm resistance to water loss.     

Focus on Ethylene: Ethylene Biosynthesis Is Promoted by Very-Long-Chain Fatty Acids during Lysigenous Aerenchyma Formation in Rice Roots

In rice (Oryza sativa) roots, lysigenous aerenchyma, which is created by programmed cell death and lysis of cortical cells, is constitutively formed under aerobic conditions, and its formation is further induced under oxygen-deficient conditions. Ethylene is involved in the induction of aerenchyma formation. reduced culm number1 (rcn1) is a rice mutant in which the gene encoding the ATP-binding cassette transporter RCN1/OsABCG5 is defective. Here, we report that the induction of aerenchyma formation was reduced in roots of rcn1 grown in stagnant deoxygenated nutrient solution (i.e. under stagnant conditions, which mimic oxygen-deficient conditions in waterlogged soils). 1-Aminocyclopropane-1-carboxylic acid synthase (ACS) is a key enzyme in ethylene biosynthesis. Stagnant conditions hardly induced the expression of ACS1 in rcn1 roots, resulting in low ethylene production in the roots. Accumulation of saturated very-long-chain fatty acids (VLCFAs) of 24, 26, and 28 carbons was reduced in rcn1 roots. Exogenously supplied VLCFA (26 carbons) increased the expression level of ACS1 and induced aerenchyma formation in rcn1 roots. Moreover, in rice lines in which the gene encoding a fatty acid elongase, CUT1-LIKE (CUT1L; a homolog of the gene encoding Arabidopsis CUT1, which is required for cuticular wax production), was silenced, both ACS1 expression and aerenchyma formation were reduced. Interestingly, the expression of ACS1, CUT1L, and RCN1/OsABCG5 was induced predominantly in the outer part of roots under stagnant conditions. These results suggest that, in rice under oxygen-deficient conditions, VLCFAs increase ethylene production by promoting 1-aminocyclopropane-1-carboxylic acid biosynthesis in the outer part of roots, which, in turn, induces aerenchyma formation in the root cortex.Aerenchyma formation is a morphological adaptation of plants to complete submergence and waterlogging of the soil, and facilitates internal gas diffusion (Armstrong, 1979; Jackson and Armstrong, 1999; Colmer, 2003; Voesenek et al., 2006; Bailey-Serres and Voesenek, 2008; Licausi and Perata, 2009; Sauter, 2013; Voesenek and Bailey-Serres, 2015). To adapt to waterlogging in soil, rice (Oryza sativa) develops lysigenous aerenchyma in shoots (Matsukura et al., 2000; Colmer and Pedersen, 2008; Steffens et al., 2011) and roots (Jackson et al., 1985b; Justin and Armstrong, 1991; Kawai et al., 1998), which is formed by programmed cell death and subsequent lysis of some cortical cells (Jackson and Armstrong, 1999; Evans, 2004; Yamauchi et al., 2013). In rice roots, lysigenous aerenchyma is constitutively formed under aerobic conditions (Jackson et al., 1985b), and its formation is further induced under oxygen-deficient conditions (Colmer et al., 2006; Shiono et al., 2011). The former and latter are designated constitutive and inducible lysigenous aerenchyma formation, respectively (Colmer and Voesenek, 2009). The gaseous plant hormone ethylene regulates adaptive growth responses of plants to submergence (Voesenek and Blom, 1989; Voesenek et al., 1993; Visser et al., 1996a,b; Lorbiecke and Sauter, 1999; Hattori et al., 2009; Steffens and Sauter, 2009; van Veen et al., 2013). Ethylene also induces lysigenous aerenchyma formation in roots of some gramineous plants (Drew et al., 2000; Shiono et al., 2008). The treatment of roots with ethylene or its precursor (1-aminocyclopropane-1-carboxylic acid [ACC]) stimulates aerenchyma formation in rice (Justin and Armstrong, 1991; Colmer et al., 2006; Yukiyoshi and Karahara, 2014), maize (Zea mays; Drew et al., 1981; Jackson et al., 1985a; Takahashi et al., 2015), and wheat (Triticum aestivum; Yamauchi et al., 2014a,b). Moreover, treatment of roots with inhibitors of ethylene action or ethylene biosynthesis effectively blocks aerenchyma formation under hypoxic conditions in maize (Drew et al., 1981; Konings, 1982; Jackson et al., 1985a; Rajhi et al., 2011).Ethylene biosynthesis is accomplished by two main successive enzymatic reactions: conversion of S-adenosyl-Met to ACC by 1-aminocyclopropane-1-carboxylic acid synthase (ACS), and conversion of ACC to ethylene by 1-aminocyclopropane-1-carboxylic acid oxidase (ACO; Yang and Hoffman, 1984). The activities of both enzymes are enhanced during aerenchyma formation under hypoxic conditions in maize root (He et al., 1996). Since the ACC content in roots of maize is increased by oxygen deficiency and is strongly correlated with ethylene production (Atwell et al., 1988), ACC biosynthesis is essential for ethylene production during aerenchyma formation in roots. In fact, exogenously supplied ACC induced ethylene production in roots of maize (Drew et al., 1979; Konings, 1982; Atwell et al., 1988) and wheat (Yamauchi et al., 2014b), even under aerobic conditions. Ethylene production in plants is inversely related to oxygen concentration (Yang and Hoffman, 1984). Under anoxic conditions, the oxidation of ACC to ethylene by ACO, which requires oxygen, is almost completely repressed (Yip et al., 1988; Tonutti and Ramina, 1991). Indeed, anoxic conditions stimulate neither ethylene production nor aerenchyma formation in maize adventitious roots (Drew et al., 1979). Therefore, it is unlikely that the root tissues forming inducible aerenchyma are anoxic, and that the ACO-mediated step is repressed. Moreover, aerenchyma is constitutively formed in rice roots even under aerobic conditions (Jackson et al., 1985b), and thus, after the onset of waterlogging, oxygen can be immediately supplied to the apical regions of roots through the constitutively formed aerenchyma.Very-long-chain fatty acids (VLCFAs; ≥20 carbons) are major constituents of sphingolipids, cuticular waxes, and suberin in plants (Franke and Schreiber, 2007; Kunst and Samuels, 2009). In addition to their structural functions, VLCFAs directly or indirectly participate in several physiological processes (Zheng et al., 2005; Reina-Pinto et al., 2009; Roudier et al., 2010; Ito et al., 2011; Nobusawa et al., 2013; Tsuda et al., 2013), including the regulation of ethylene biosynthesis (Qin et al., 2007). During fiber cell elongation in cotton ovules, ethylene biosynthesis is enhanced by treatment with saturated VLCFAs, especially 24-carbon fatty acids, and is suppressed by an inhibitor of VLCFA biosynthesis (Qin et al., 2007). The first rate-limiting step in VLCFA biosynthesis is condensation of acyl-CoA with malonyl-CoA by β-ketoacyl-CoA synthase (KCS; Joubès et al., 2008). KCS enzymes are thought to determine the substrate and tissue specificities of fatty acid elongation (Joubès et al., 2008). The Arabidopsis (Arabidopsis thaliana) genome has 21 KCS genes (Joubès et al., 2008). In the Arabidopsis cut1 mutant, which has a defect in the gene encoding CUT1 that is required for cuticular wax production (i.e. one of the KCS genes), the expression of AtACO genes and growth of root cells were reduced when compared with the wild type (Qin et al., 2007). Furthermore, expression of the AtACO genes was rescued by exogenously supplied saturated VLCFAs (Qin et al., 2007). These observations imply that VLCFAs or their derivatives work as regulatory factors for gene expression during some physiological processes in plants.reduced culm number1 (rcn1) was first identified as a rice mutant with a low tillering rate in a paddy field (Takamure and Kinoshita, 1985; Yasuno et al., 2007). The rcn1 (rcn1-2) mutant has a single nucleotide substitution in the gene encoding a member of the ATP-binding cassette (ABC) transporter subfamily G, RCN1/OsABCG5, causing an Ala-684Pro substitution (Yasuno et al., 2009). The mutation results in several mutant phenotypes, although the substrates of RCN1/OsABCG5 have not been determined (Ureshi et al., 2012; Funabiki et al., 2013; Matsuda et al., 2014). We previously found that the rcn1 mutant has abnormal root morphology, such as shorter root length and brownish appearance of roots, under stagnant (deoxygenated) conditions (which mimics oxygen-deficient conditions in waterlogged soils). We also found that the rcn1 mutant accumulates less of the major suberin monomers originating from VLCFAs in the outer part of adventitious roots, and this results in a reduction of a functional apoplastic barrier in the root hypodermis (Shiono et al., 2014a).The objective of this study was to elucidate the molecular basis of inducible aerenchyma formation. To this end, we examined lysigenous aerenchyma formation and ACC, ethylene, and VLCFA accumulation and their biosyntheses in rcn1 roots. Based on the results of these studies, we propose that VLCFAs are involved in inducible aerenchyma formation through the enhancement of ethylene biosynthesis in rice roots.     

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

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

Male Sterile2 encodes a plastid-localized fatty acyl carrier protein reductase required for pollen exine development in Arabidopsis

Male Sterile2 (MS2) is predicted to encode a fatty acid reductase required for pollen wall development in Arabidopsis (Arabidopsis thaliana). Transient expression of MS2 in tobacco (Nicotiana benthamiana) leaves resulted in the accumulation of significant levels of C16 and C18 fatty alcohols. Expression of MS2 fused with green fluorescent protein revealed that an amino-terminal transit peptide targets the MS2 to plastids. The plastidial localization of MS2 is biologically important because genetic complementation of MS2 in ms2 homozygous plants was dependent on the presence of its amino-terminal transit peptide or that of the Rubisco small subunit protein amino-terminal transit peptide. In addition, two domains, NAD(P)H-binding domain and sterile domain, conserved in MS2 and its homologs were also shown to be essential for MS2 function in pollen exine development by genetic complementation testing. Direct biochemical analysis revealed that purified recombinant MS2 enzyme is able to convert palmitoyl-Acyl Carrier Protein to the corresponding C16:0 alcohol with NAD(P)H as the preferred electron donor. Using optimized reaction conditions (i.e. at pH 6.0 and 30°C), MS2 exhibits a K_m for 16:0-Acyl Carrier Protein of 23.3 ± 4.0 μm, a V_max of 38.3 ± 4.5 nmol mg⁻¹ min⁻¹, and a catalytic efficiency/K_m of 1,873 m⁻¹ s⁻¹. Based on the high homology of MS2 to other characterized fatty acid reductases, it was surprising that MS2 showed no activity against palmitoyl- or other acyl-coenzyme A; however, this is consistent with its plastidial localization. In summary, genetic and biochemical evidence demonstrate an MS2-mediated conserved plastidial pathway for the production of fatty alcohols that are essential for pollen wall biosynthesis in Arabidopsis.In flowering plants, the life cycle alternates between diploid sporophyte and haploid gametophyte generations. Pollen grains play a biologically protective role for the haploid male sperm cells surrounded by the outer cell wall lipidic biopolymers called the exine (Blackmore et al., 2007; Li and Zhang, 2010; Ariizumi and Toriyama, 2011). Pollen exine protects the gametophyte against pathogen attack, dehydration, and UV irradiation as well as facilitates the pollination process, including pollen recognition and adhesion to the stigma. The highly durable exine that occurs throughout flowering plants is thought to play an essential role in land colonization by plants (Chaloner, 1976).The exine is mainly composed of the biopolymer sporopollenin and contains two sublayers, the sexine and nexine (Zinkl et al., 1999). The biochemical nature of pollen exine remains largely unknown because of the technical difficulties in purifying and obtaining large quantities of materials for analysis. In addition, sporopollenin is highly insoluble, resistant to degradation, and exceptionally stable (Brooks and Shaw, 1968; Bubert et al., 2002). Current evidence suggests that the major components of sporopollenin are derivatives of aliphatics, such as fatty acids and phenolic compounds (Bubert et al., 2002; Blackmore et al., 2007).Tapetum, the innermost sporophytic anther wall layer, is thought to play a major role in actively synthesizing and secreting sporopollenin precursors onto the microspore surface for pollen exine polymerization and patterning (Bedinger, 1992; Li and Zhang, 2010; Ariizumi and Toriyama, 2011). The model dicot plant Arabidopsis (Arabidopsis thaliana) and many other plants have a secretory-type tapetum with specialized structures such as tapetosomes in the cells, which accumulate lipidic components (Huysmans et al., 1998). The outer surface of Arabidopsis pollen grains displays elegant reticulate cavities with abundant pollen coat (tryphine) deposited inside the pollen exine.Pollen exine patterning appears to include at least three major developmental events: callose wall formation, primexine formation, and sporopollenin synthesis (Ariizumi and Toriyama, 2011). Exine formation commences after meiosis, with the accumulation of lipidic precursors onto the primexine surrounding newly formed microspores between the callose wall and the microspore plasma membrane (Paxson-Sowders et al., 2001; Blackmore et al., 2007). After the first pollen mitosis, the synthesis of the exine is almost complete; and during later stages of pollen exine formation, the pectocellulosic intine and the tryphine, called the pollen coat, are deposited onto the pollen wall (Piffanelli et al., 1998). Recent genetic and biochemical investigations showed that some genes, including MALE STERILITY1 (MS1), MS2, CER1, NO EXINE FORMATION1, FACELESS POLLEN1, CYP703A2, ACYL-COA SYNTHETASE5 (ACOS5), CYP704B1, TETRAKETIDE α-PYRONE REDUCTASE1 and -2 (TKPR1/2), LAP6/POLYKETIDE SYNTHASE A (PKSA), and LAP5/POLYKETIDE SYNTHASE B (PKSB) in Arabidopsis (Aarts et al., 1995, 1997; Wilson et al., 2001; Ariizumi et al., 2003, 2004; Morant et al., 2007; de Azevedo Souza et al., 2009; Dobritsa et al., 2009, 2010; Grienenberger et al., 2010; Kim et al., 2010b) as well as Tapetum Degeneration Retardation, Wax-Deficient Anther1, CYP704B2, C6, Postmeiotic Deficient Anther1, and Persistent Tapetal Cell1 in rice (Oryza sativa; Jung et al., 2006; Zhang et al., 2008, 2010, 2011; Hu et al., 2010; Li et al., 2006, 2010, 2011; Li and Zhang, 2010), are required for pollen exine synthesis. However, relatively little has been described on the biochemical aspects of these gene products.Fatty alcohols are widely observed in plants, animals, and algae in free forms (the component of cuticular lipids) but more frequently in esterified (wax esters) or etherified (glyceryl ethers) forms. Fatty alcohols and their derivatives are major components of the lipidic anther cuticle and pollen wall (Ahlers et al., 1999; Kunst and Samuels, 2003; Jung et al., 2006; Li et al., 2010). Previous investigations revealed that fatty acyl-CoAs are thought to be used as substrates for the production of fatty alcohols by fatty acyl-coenzyme A reductase (FAR) in garden pea (Pisum sativum), jojoba (Simmondsia chinensis), Arabidopsis, wheat (Triticum aestivum), mouse (Mus musculus), and silk moth (Bombyx mori; Aarts et al., 1997; Metz et al., 2000; Wang et al., 2002; Moto et al., 2003; Cheng and Russell, 2004; Rowland et al., 2006; Doan et al., 2009; Domergue et al., 2010).MS2 was assumed to encode a FAR-like protein that converts fatty acids to alcohols. MS2 was shown to be expressed in the tapetum shortly after the microspore was released from the tetrad (Aarts et al., 1997). ms2 mutants display abnormal pollen wall development, which is sensitive to acetolysis treatment, causing reduced pollen fertility (Aarts et al., 1997; Dobritsa et al., 2009). However, detailed biochemical characterization of the MS2 enzyme has not been performed. Recently, recombinant bacteria expressing five Arabidopsis FAR homologs were shown to produce fatty alcohols with carbon lengths of C14, C16, and C18 from endogenous bacterial fatty acids. Bacteria expressing MS2 are able to form C14:0, C16:0, and C18:1 alcohols (Doan et al., 2009). Furthermore, yeast cells expressing FAR1, FAR4, and FAR5 are able to produce alcohols using distinct but overlapping substrates with a chain length ranging from C18:0 to C24:0 (Domergue et al., 2010).In this study, we report the biochemical characterization of MS2. We show that MS2 encodes a fatty acyl-Acyl Carrier Protein (ACP) reductase, and the purified recombinant MS2 enzyme from Escherichia coli is able to convert the preferred substrate palmitoyl-ACP to C16:0 alcohol in the presence of NAD(P)H. In addition, MS2 possesses an N-terminal transit peptide that is necessary for localization to the plastid. The biological significance of MS2 subcellular localization, and the presence of conserved domains/motifs within MS2, were demonstrated by genetic complementation of ms2 mutants. This work, therefore, demonstrates the involvement of the plastid in primary fatty alcohol synthesis required for pollen wall development in Arabidopsis.     

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.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

A Host RNA Helicase-Like Protein,AtRH8, Interacts with the Potyviral Genome-Linked Protein,VPg, Associates with the Virus Accumulation Complex,and Is Essential for Infection

The viral genome-linked protein, VPg, of potyviruses is a multifunctional protein involved in viral genome translation and replication. Previous studies have shown that both eukaryotic translation initiation factor 4E (eIF4E) and eIF4G or their respective isoforms from the eIF4F complex, which modulates the initiation of protein translation, selectively interact with VPg and are required for potyvirus infection. Here, we report the identification of two DEAD-box RNA helicase-like proteins, PpDDXL and AtRH8 from peach (Prunus persica) and Arabidopsis (Arabidopsis thaliana), respectively, both interacting with VPg. We show that AtRH8 is dispensable for plant growth and development but necessary for potyvirus infection. In potyvirus-infected Nicotiana benthamiana leaf tissues, AtRH8 colocalizes with the chloroplast-bound virus accumulation vesicles, suggesting a possible role of AtRH8 in viral genome translation and replication. Deletion analyses of AtRH8 have identified the VPg-binding region. Comparison of this region and the corresponding region of PpDDXL suggests that they are highly conserved and share the same secondary structure. Moreover, overexpression of the VPg-binding region from either AtRH8 or PpDDXL suppresses potyvirus accumulation in infected N. benthamiana leaf tissues. Taken together, these data demonstrate that AtRH8, interacting with VPg, is a host factor required for the potyvirus infection process and that both AtRH8 and PpDDXL may be manipulated for the development of genetic resistance against potyvirus infections.Plant viruses are obligate intracellular parasites that infect many agriculturally important crops and cause severe losses each year. One of the common characteristics of plant viruses is their relatively small genome that encodes a limited number of viral proteins, making them dependent on host factors to fulfill their infection cycles (Maule et al., 2002; Whitham and Wang, 2004; Nelson and Citovsky, 2005; Decroocq et al., 2006). In order to establish a successful infection, the invading virus must recruit an array of host proteins (host factors) to translate and replicate its genome and to move locally from cell to cell via the plasmodesmata and systemically via the vascular system. It has been suggested that down-regulation or mutation of some of the required host factors may result in recessively inherited resistance to viruses (Kang et al., 2005b).Potyviruses, belonging to the genus Potyvirus in the family Potyviradae, constitute the largest group of plant viruses (Rajamäki et al., 2004). Potyviruses have a single positive-strand RNA genome approximately 10 kb in length, with a viral genome-linked protein (VPg) covalently attached to the 5′ end and a poly(A) tail at the 3′ end (Urcuqui-Inchima et al., 2001; Rajamäki et al., 2004). The viral genome contains a single open reading frame (ORF) that translates into a polypeptide with a molecular mass of approximately 350 kD, which is cleaved into 10 mature proteins by viral proteases (Urcuqui-Inchima et al., 2001). Recently, a novel viral protein resulting from a frameshift in the P3 cistron has been reported (Chung et al., 2008). Of the 11 viral proteins, VPg is a multifunctional protein and the only other viral protein present in the viral particles (virions) besides the coat protein and the cylindrical inclusion protein (CI; Oruetxebarria et al., 2001; Puustinen et al., 2002; Gabrenaite-Verkhovskaya et al., 2008). The nonstructural protein is linked to the viral RNA by a phosphodiester bond between the 5′ terminal uridine residue of the RNA and the O⁴-hydroxyl group of amino acid Tyr (Murphy et al., 1996; Oruetxebarria et al., 2001; Puustinen et al., 2002). Mutation of the Tyr residue that links VPg to the viral RNA abolishes virus infectivity completely (Murphy et al., 1996). In infected cells, VPg and its precursor NIa are present in the nucleus and in the membrane-associated virus replication vesicles in the cytoplasm (Carrington et al., 1993; Rajamäki and Valkonen, 2003; Cotton et al., 2009). As a component of the replication complex, VPg may serve as a primer for viral RNA replication (Puustinen and Mäkinen, 2004) and as an analog of the m⁷G cap of mRNAs for the viral genome to recruit the translation complex for translation (Michon et al., 2006; Beauchemin et al., 2007; Khan et al., 2008). Furthermore, VPg has been suggested to be an avirulence factor for recessive resistance genes in diverse plant species (Moury et al., 2004; Kang et al., 2005b; Bruun-Rasmussen et al., 2007). Thus, VPg plays a pivotal role in the virus infection process. The molecular identification of VPg-interacting host proteins and the subsequent functional characterization of such interactions may advance knowledge of the intricate virus replication mechanisms and help develop novel antiviral strategies.Previous studies have shown that VPg and its precursor NIa interact with several host proteins, including three essential components of the host protein translation apparatus (Thivierge et al., 2008). The first protein is the cellular translation initiation factor eIF4E or its isoform eIF(iso)4E, identified through a yeast two-hybrid screen using VPg as a bait (Wittmann et al., 1997; Schaad et al., 2000). The protein complex of VPg and eIF4E is an essential component for virus infectivity (Robaglia and Caranta, 2006). Mutations and knockout of eIF4E or eIF(iso)4E confer resistance to infection (Lellis et al., 2002; Ruffel et al., 2002; Nicaise et al., 2003; Gao et al., 2004; Kang et al., 2005a; Ruffel et al., 2005; Decroocq et al., 2006; Bruun-Rasmussen et al., 2007). It is well known that potyviruses recruit selectively one of the eIF4E isoforms, depending on specific virus-host combinations (German-Retana et al., 2008). For instance, in Arabidopsis (Arabidopsis thaliana), eIF(iso)4E is required for infection by Turnip mosaic virus (TuMV), Plum pox virus (PPV), and Lettuce mosaic virus, while eIF4E is indispensable for infection by Clover yellow vein virus (Duprat et al., 2002; Lellis et al., 2002; Sato et al., 2005; Decroocq et al., 2006). The second cellular protein interacting with VPg is another translation initiation factor, eIF4G. Analysis of Arabidopsis knockout mutants for eIF4G or its isomers eIF(iso)4G1 and eIF(iso)4G2 has yielded results supporting the idea that the recruitment of eIF4G for potyvirus infection is also isoform dependent (Nicaise et al., 2007). Recently, poly(A)-binding protein (PABP), the translation initiation factor that bridges the 5′ and 3′ termini of the mRNA into proximity, has been proposed to be essential for efficient multiplication of TuMV (Dufresne et al., 2008). PABP was previously documented to interact with NIa, a VPg precursor containing both VPg and the proteinase NIa-Pro (Léonard et al., 2004). As the translation factors eIF(iso)4E and PABP have been found to be internalized in virus-induced vesicles, it has been suggested that the interactions between VPg and these translation factors are crucial for viral RNA translation and/or replication (Beauchemin and Laliberté, 2007; Beauchemin et al., 2007; Cotton et al., 2009). Besides these three translation factors, a Cys-rich plant protein, potyvirus VPg-interaction protein, was also found to associate with VPg (Dunoyer et al., 2004). This plant-specific VPg-interacting host protein contains a PHD finger domain and acts as an ancillary factor to support potyvirus infection and movement (Dunoyer et al., 2004).In this study, we describe the identification of an Arabidopsis DEAD-box RNA helicase (DDX), AtRH8, and a peach (Prunus persica) DDX-like protein, PpDDXL, both interacting with the potyviral VPg protein. Using the atrh8 mutant, we demonstrate that AtRH8 is not required for plant growth and development in Arabidopsis but is necessary for infection by two plant potyviruses, PPV and TuMV. Furthermore, we present evidence that AtRH8 colocalizes with the virus accumulation complex in potyvirus-infected leaf tissues, which reveals a possible role of AtRH8 in virus infection. Finally, we have identified the VPg-binding region (VPg-BR) of AtRH8 and PpDDX and show that overexpression of the VPg-BR either from AtRH8 or PpDDXL suppresses virus accumulation.