Microalgae Synthesize Hydrocarbons from Long-Chain Fatty Acids via a Light-Dependent Pathway

Microalgae are considered a promising platform for the production of lipid-based biofuels. While oil accumulation pathways are intensively researched, the possible existence of a microalgal pathways converting fatty acids into alka(e)nes has received little attention. Here, we provide evidence that such a pathway occurs in several microalgal species from the green and the red lineages. In Chlamydomonas reinhardtii (Chlorophyceae), a C17 alkene, n-heptadecene, was detected in the cell pellet and the headspace of liquid cultures. The Chlamydomonas alkene was identified as 7-heptadecene, an isomer likely formed by decarboxylation of cis-vaccenic acid. Accordingly, incubation of intact Chlamydomonas cells with per-deuterated D₃₁-16:0 (palmitic) acid yielded D₃₁-18:0 (stearic) acid, D₂₉-18:1 (oleic and cis-vaccenic) acids, and D₂₉-heptadecene. These findings showed that loss of the carboxyl group of a C₁₈ monounsaturated fatty acid lead to heptadecene formation. Amount of 7-heptadecene varied with growth phase and temperature and was strictly dependent on light but was not affected by an inhibitor of photosystem II. Cell fractionation showed that approximately 80% of the alkene is localized in the chloroplast. Heptadecane, pentadecane, as well as 7- and 8-heptadecene were detected in Chlorella variabilis NC64A (Trebouxiophyceae) and several Nannochloropsis species (Eustigmatophyceae). In contrast, Ostreococcus tauri (Mamiellophyceae) and the diatom Phaeodactylum tricornutum produced C₂₁ hexaene, without detectable C₁₅-C₁₉ hydrocarbons. Interestingly, no homologs of known hydrocarbon biosynthesis genes were found in the Nannochloropsis, Chlorella, or Chlamydomonas genomes. This work thus demonstrates that microalgae have the ability to convert C₁₆ and C₁₈ fatty acids into alka(e)nes by a new, light-dependent pathway.Hydrocarbons derived from fatty acids (i.e. alkanes and alkenes) are ubiquitous in plant and insect outermost tissues where they often represent a major part of cuticular waxes and play an essential role in preventing water loss from the organisms to the dry terrestrial environment (Hadley, 1989; Kunst et al., 2005). In several insect species, select cuticular alkenes also act as sex pheromones (Wicker-Thomas and Chertemps, 2010). Occurrence of alkanes or alkenes has also been reported in various microorganisms (Ladygina et al., 2006; Wang and Lu, 2013). For example, synthesis of hydrocarbons is widespread in cyanobacteria (Coates et al., 2014), and it is thought that cyanobacterial alka(e)nes contribute significantly to the hydrocarbon cycle of the upper ocean (Lea-Smith et al., 2015).Alka(e)nes of various chain lengths are important targets for biotechnology because they are major components of gasoline (mainly C5-C9 hydrocarbons), jet fuels (C5-C16), and diesel fuels (C12-C20). The alkane biosynthetic pathway of plants has been partly elucidated (Lee and Suh, 2013; Bernard and Joubès, 2013), but the use of plant hydrocarbons as a renewable source of liquid fuels is hampered by predominance of constituents with high carbon numbers (>C25), which entails solid state at ambient temperature (Jetter and Kunst, 2008). Therefore, there is great interest in the microbial pathways of hydrocarbon synthesis producing shorter chain compounds (C15-C19). In cyanobacteria, hydrocarbons are produced by two distinct pathways. The first one comprises the sequential action of an acyl-ACP reductase transforming a C15-C19 fatty acyl-ACP into a fatty aldehyde and an aldehyde-deformylating oxygenase catalyzing the oxidative cleavage of the fatty aldehyde into alka(e)ne and formic acid (Schirmer et al., 2010; Li et al., 2012). The second pathway involves a type I polyketide synthase that elongates and decarboxylates fatty acids to form alkenes with a terminal double bond (Mendez-Perez et al., 2011). Additional pathways of alkene synthesis have been described in bacteria. In Micrococcus luteus, a three-gene cluster has been shown to control the head-to-head condensation of fatty acids to form very-long-chain alkenes with internal double bonds (Beller et al., 2010). Direct decarboxylation of a long-chain fatty acid into a terminal alkene has also been reported and is catalyzed by a cytochrome P450 in Jeotgalicoccus spp. (Rude et al., 2011) and by a nonheme iron oxidase in Pseudomonas (Rui et al., 2014).Among microbes, microalgae would be ideally suited to harness the synthesis of hydrocarbons from fatty acid precursors because they are photosynthetic organisms combining a high biomass productivity (León-Bañares et al., 2004; Beer et al., 2009; Malcata, 2011; Wijffels et al., 2013) and a strong capacity to synthesize and accumulate fatty acids (Hu et al., 2008; Harwood and Guschina, 2009; Liu and Benning 2013). However, studies on microalgal alka(e)ne synthesis are scarce. In some diatoms and other algal species, a very-long-chain alkene, a C21 hexaene, has been found (Lee et al., 1970; Lee and Loeblich, 1971). Other very-long-chain alkenes have been described in the slow-growing colonial Chlorophycea Botryococcus brauni, which excretes a variety of hydrophobic compounds including C27 n-alkadienes (Metzger and Largeau, 2005; Jin et al., 2016). A decarbonylase activity converting a fatty n-aldehyde substrate to a n-alkane has been shown in B. brauni (Dennis and Kolattukudy, 1992); however, the corresponding protein has not been identified so far. Presence of shorter chain alka(e)nes in some microalga species has been reported in the context of geochemical studies (Han et al., 1968; Gelpi et al., 1970; Tornabene et al., 1980; Afi et al., 1996) but the biology of these compounds has not been investigated further.Here, we show that alka(e)nes with C15 to C17 chains can be detected in several model microalgae and originate from fatty acid metabolism. In Chlamydomonas reinhardtii and Chlorella variabilis NC64A, 7-heptadecene is identified as the major hydrocarbon produced, and we demonstrate that its synthesis depends strictly on light and uses cis-vaccenic acid as a precursor. We also show the presence of C15 to C17 alka(e)nes in Nannochloropsis sp., a model microalga from the red lineage. Absence of homologs to known hydrocarbon synthesis genes in the genomes of Chlamydomonas, Chlorella, and Nannochloropsis indicates that a hitherto unknown type of alka(e)ne-producing pathway operates in these microalgae. The wide occurrence of microalgae in marine environments suggests that microalgal alka(e)nes contribute significantly to the ocean’s hydrocarbon cycle.     

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.     

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.     

Arabidopsis ECERIFERUM2 Is a Component of the Fatty Acid Elongation Machinery Required for Fatty Acid Extension to Exceptional Lengths

Primary aerial surfaces of land plants are coated by a lipidic cuticle, which forms a barrier against transpirational water loss and protects the plant from diverse stresses. Four enzymes of a fatty acid elongase complex are required for the synthesis of very-long-chain fatty acid (VLCFA) precursors of cuticular waxes. Fatty acid elongase substrate specificity is determined by a condensing enzyme that catalyzes the first reaction carried out by the complex. In Arabidopsis (Arabidopsis thaliana), characterized condensing enzymes involved in wax synthesis can only elongate VLCFAs up to 28 carbons (C28) in length, despite the predominance of C29 to C31 monomers in Arabidopsis stem wax. This suggests additional proteins are required for elongation beyond C28. The wax-deficient mutant eceriferum2 (cer2) lacks waxes longer than C28, implying that CER2, a putative BAHD acyltransferase, is required for C28 elongation. Here, we characterize the cer2 mutant and demonstrate that green fluorescent protein-tagged CER2 localizes to the endoplasmic reticulum, the site of VLCFA biosynthesis. We use site-directed mutagenesis to show that the classification of CER2 as a BAHD acyltransferase based on sequence homology does not fit with CER2 catalytic activity. Finally, we provide evidence for the function of CER2 in C28 elongation by an assay in yeast (Saccharomyces cerevisiae).Land plants have a lipidic cuticle that seals the outer surface of all of their primary aerial organs. Structurally, the cuticle consists of two components, cutin and cuticular waxes. Together these form a hydrophobic barrier that plays a critical role in plant survival by restricting nonstomatal water loss (Riederer and Schreiber, 2001). Cuticles also protect the plant from biotic and abiotic stresses, profoundly affect plant-insect interactions (Müller, 2006), prevent epidermal fusions (Sieber et al., 2000), and are involved in drought stress signaling (Wang et al., 2011).Cutin is a polymer of mainly midchain- and ω-hydroxy and -epoxy 16 carbon (C16) and C18 fatty acids, which are cross-linked in ester bonds directly or through a glycerol backbone (Pollard et al., 2008). Cuticular waxes are aliphatic monomers that are deposited within the cutin matrix as intracuticular wax, and on top of it as epicuticular wax film and crystals. Wax is a heterogeneous mixture of very-long-chain fatty acids (VLCFAs) and their alkane, aldehyde, alcohol, ketone, and ester derivatives, which typically range from C24 to C32 in length (Samuels et al., 2008). The composition of cuticular wax varies greatly among species and tissues, often providing physical and chemical properties to the plant surface that are advantageous in specific environments.Genetic analyses have revealed that a fatty acid elongase (FAE) complex is responsible for the synthesis of VLCFA wax precursors (Millar et al., 1999; Fiebig et al., 2000; Kunst and Samuels, 2009). FAE complexes are heterotetramers of independently transcribed, monofunctional proteins localized to the endoplasmic reticulum (ER). Together, they catalyze a series of four reactions to elongate long-chain acyl-CoAs or very-long-chain acyl-CoAs by sequential addition of two carbon units. The condensing enzyme, or β-ketoacyl-CoA synthase (KCS), catalyzes the first reaction in this sequence and is both rate limiting and specific for the chain length of acyl-CoA synthesized (Millar and Kunst, 1997). Two very dissimilar families of KCSs have been identified in Arabidopsis (Arabidopsis thaliana): a FAE1-type family homologous to the first such KCS enzyme discovered in association with seed oil biosynthesis (Kunst et al., 1992; James et al., 1995; Lassner et al., 1996), and an ELONGATION DEFECTIVE (ELO)-like family homologous to the yeast (Saccharomyces cerevisiae) ELO family responsible for sphingolipid synthesis (Dunn et al., 2004). To date, no function has been ascribed to Arabidopsis ELOs. Of the 21 FAE1-type KCS enzymes in Arabidopsis (Joubès et al., 2008), 11 have been shown by microarray analysis to be up-regulated in the stem epidermis (Suh et al., 2005). Only one of these, ECERIFERUM6 (CER6/KCS6/CUT1; Millar et al., 1999; Fiebig et al., 2000; Joubès et al., 2008), has a dominant role in the elongation of VLCFAs for cuticular wax synthesis, as CER6 suppression results in a dramatic reduction of all wax monomers longer than C24 (Millar et al., 1999). Heterologous expression of CER6 in yeast has demonstrated that the CER6 condensing enzyme can produce C28 VLCFAs (O. Rowland and L. Kunst, unpublished data). However, CER6 appears to be unable to produce VLCFAs longer than C28 in yeast; this presents a problem as the bulk of Arabidopsis stem wax is made up of C29 alkanes, secondary alcohols, and ketones derived from C30 VLCFAs. Mutant screens have not revealed any other KCS enzymes necessary for VLCFA elongation past C28 in Arabidopsis. Therefore, there may be other proteins unrelated to condensing enzymes that are required for acyl chain extension beyond C28 that remain unknown.The wax-deficient mutant cer2 shows a dramatic reduction in all stem waxes longer than C28 and increased accumulation of waxes C28 or shorter, suggesting that CER2 has a role in the final steps of VLCFA elongation. Surprisingly, the cer2 mutation has been mapped to At4g24510 (Negruk et al., 1996; Xia et al., 1996), a gene homologous to plant BAHD acyltransferases. However, the CER2 protein was reported to localize exclusively to the nucleus (Xia et al., 1997). This does not fit with CER2 annotation as a BAHD acyltransferase, as all characterized BAHD acyltransferases are soluble cytosolic enzymes (D’Auria, 2006).The objective of this work was to more precisely evaluate the role of CER2 in fatty acid elongation using a new CER2 allele, cer2-5 (Columbia-0 [Col-0] ecotype). We provide evidence that CER2 has a metabolic function specific to wax synthesis, and that the CER2 homolog CER2-LIKE1 has an analogous role in leaf wax synthesis. Despite the classification of CER2 as a BAHD acyltransferase based on sequence homology, we demonstrate that CER2 cannot share the catalytic mechanism that has been confirmed for other members of the BAHD family, and provide biochemical support for a function of CER2 in VLCFA elongation by an assay in yeast.     

ECERIFERUM2-LIKE Proteins Have Unique Biochemical and Physiological Functions in Very-Long-Chain Fatty Acid Elongation

The extension of very-long-chain fatty acids (VLCFAs) for the synthesis of specialized apoplastic lipids requires unique biochemical machinery. Condensing enzymes catalyze the first reaction in fatty acid elongation and determine the chain length of fatty acids accepted and produced by the fatty acid elongation complex. Although necessary for the elongation of all VLCFAs, known condensing enzymes cannot efficiently synthesize VLCFAs longer than 28 carbons, despite the prevalence of C28 to C34 acyl lipids in cuticular wax and the pollen coat. The eceriferum2 (cer2) mutant of Arabidopsis (Arabidopsis thaliana) was previously shown to have a specific deficiency in cuticular waxes longer than 28 carbons, and heterologous expression of CER2 in yeast (Saccharomyces cerevisiae) demonstrated that it can modify the acyl chain length produced by a condensing enzyme from 28 to 30 carbon atoms. Here, we report the physiological functions and biochemical specificities of the CER2 homologs CER2-LIKE1 and CER2-LIKE2 by mutant analysis and heterologous expression in yeast. We demonstrate that all three CER2-LIKEs function with the same small subset of condensing enzymes, and that they have different effects on the substrate specificity of the same condensing enzyme. Finally, we show that the changes in acyl chain length caused by each CER2-LIKE protein are of substantial importance for cuticle formation and pollen coat function.The extension of fatty acids to lengths greater than 28 carbons (C28) is an exceptional process in plant metabolism in that it requires unique biochemical machinery, and the elongation products are used for the synthesis of specialized plant metabolites. Derivatives of C30 to C34 fatty acids make up the bulk of plant cuticular wax, which coats all of a plant’s primary aerial surfaces. Cuticular wax serves as a barrier against transpirational water loss (Riederer and Schreiber, 2001) and protects the plant from both biotic (Eigenbrode, 1996) and abiotic (Grace and van Gardingen, 1996) stresses. C30 to C34 fatty acid-derived lipids are also components of the pollen coat, where they function in pollen hydration and germination on dry stigma (Elleman et al., 1992; Preuss et al., 1993).The core complex that elongates long-chain fatty acids (C16–C18) to very-long-chain fatty acids (VLCFAs; C20–C34) consists of four interacting proteins localized to the endoplasmic reticulum (ER). β-Keto-acyl-CoA synthases (KCSs), also known as condensing enzymes, catalyze the first reaction required for VLCFA elongation, condensing malonyl-CoA with an acyl-CoA (n) to produce a β-keto-acyl-CoA (n + 2). Condensation is both a specific and rate-limiting step in elongation (Millar and Kunst, 1997). Chain length specificity of KCSs is of particular importance because VLCFA length determines the downstream use of the fatty acid (for review, see Joubès et al., 2008; Haslam and Kunst, 2013a). There are two families of condensing enzymes in Arabidopsis (Arabidopsis thaliana). The ELONGATION-DEFECTIVE (ELO)-LIKE family is homologous to yeast (Saccharomyces cerevisiae) ELOs, and has putative functions in sphingolipid biosynthesis (Quist et al., 2009). Although our current understanding of plant ELO-LIKE physiological function and biochemical activity is limited, the mechanism of yeast Elo protein activity has been thoroughly investigated (Denic and Weissman, 2007). The FATTY ACID ELONGATION1 (FAE1)-type family is homologous to the first condensing enzyme identified in Arabidopsis, which is required for the synthesis of C20 to C22 VLCFAs in Arabidopsis oilseeds. Many of the 21 FAE1-type condensing enzymes of Arabidopsis have been characterized using reverse genetics and heterologous expression in yeast (Trenkamp et al., 2004; Blacklock and Jaworski, 2006; Paul et al., 2006; Tresch et al., 2012). This work has revealed the intriguing caveat that, although FAE1-type KCSs are involved in the synthesis of diverse downstream metabolites and use a broad range of acyl chain lengths, none are able to efficiently elongate VLCFAs beyond C28 (for review, see Haslam and Kunst, 2013a), which is essential for the production of cuticular wax components.Eceriferum2 (cer2) and glossy2 (gl2) mutants of Arabidopsis and Zea mays, respectively, are deficient in specific VLCFA-derived waxes longer than C28 (Bianchi et al., 1975; McNevin et al., 1993; Jenks et al., 1995). Both mutations were mapped to genes that do not resemble any component of the elongase complex (Tacke et al., 1995; Xia et al., 1996), but are homologous to the BAHD family of acyltransferases (St-Pierre et al., 1998). However, site-directed mutagenesis of conserved acyltransferase catalytic site amino acids in CER2 revealed that this motif is not required for CER2 function in cuticular wax synthesis (Haslam et al., 2012).CER6 is a condensing enzyme necessary for the accumulation of stem cuticular waxes in Arabidopsis, but when expressed in yeast, CER6 can only elongate VLCFAs to C28. When CER2 is expressed in yeast, it has no elongation activity. However, coexpression of CER2 and CER6 results in efficient production of C30 VLCFAs. Coexpression of CER2 with LfKCS45, a condensing enzyme from the crucifer Lesquerella fendleri that generates C28 and a small amount of C30 VLCFAs (Moon et al., 2004), does not alter product chain length (Haslam et al., 2012). Based on these observations, it was hypothesized that CER2 modifies the chain length specificity of the core elongase complex by interaction with specific KCS enzymes (Haslam et al., 2012).CER2 homologs are found in diverse flowering plant lineages, and many species have multiple CER2 homologs (Tuominen et al., 2011). A BLAST search of proteins from Arabidopsis identified two sequences with substantial similarity to CER2. {"type":"entrez-protein","attrs":{"text":"NP_193120","term_id":"15236357","term_text":"NP_193120"}}NP_193120 is 36% identical to CER2, and is encoded by the gene At4g13840. We named this gene CER2-LIKE1 (also known as CER26) (Pascal et al., 2013). {"type":"entrez-protein","attrs":{"text":"NP_566741","term_id":"18403923","term_text":"NP_566741"}}NP_566741 is 38% identical to CER2, and is encoded by the gene At3g23840. We named this gene CER2-LIKE2 (also named CER26-LIKE) (Pascal et al., 2013). Characterization of a cer2-like1 null mutant revealed a role for the CER2-LIKE1 protein in the elongation of leaf wax precursors beyond C30, analogous to the role of CER2 in C28 elongation in stems (Haslam et al., 2012; Pascal et al., 2013). cer2 cer2-like1 double mutants are deficient in the formation of wax components longer than C28 in both stems and leaves. As the cer2 single mutant has no leaf wax phenotype, the additive effect of these two mutations on leaf wax composition indicates that there is partial functional redundancy between the two genes.A comprehensive investigation of the biochemical and physiological functions of CER2-LIKE proteins is necessary. Beyond the value of knowing the specific roles of each homolog, such an investigation has potential to elucidate the nature of CER2-LIKE protein function. With this objective, we used our data to address the following questions: (1) Do CER2-LIKE proteins function with CER6 alone, or can they modify the activity of other FAE1-type condensing enzymes? (2) Do CER2-LIKE proteins have different effects on the substrate specificity of the same condensing enzyme, or is substrate specificity determined exclusively by the condensing enzyme? (3) What is the physiological relevance of the subtle changes in acyl lipid chain length that CER2-LIKE proteins induce?     

Peroxisomal ATP Import Is Essential for Seedling Development in Arabidopsis thaliana

Several recent proteomic studies of plant peroxisomes indicate that the peroxisomal matrix harbors multiple ATP-dependent enzymes and chaperones. However, it is unknown whether plant peroxisomes are able to produce ATP by substrate-level phosphorylation or whether external ATP fuels the energy-dependent reactions within peroxisomes. The existence of transport proteins that supply plant peroxisomes with energy for fatty acid oxidation and other ATP-dependent processes has not previously been demonstrated. Here, we describe two Arabidopsis thaliana genes that encode peroxisomal adenine nucleotide carriers, PNC1 and PNC2. Both proteins, when fused to enhanced yellow fluorescent protein, are targeted to peroxisomes. Complementation of a yeast mutant deficient in peroxisomal ATP import and in vitro transport assays using recombinant transporter proteins revealed that PNC1 and PNC2 catalyze the counterexchange of ATP with ADP or AMP. Transgenic Arabidopsis lines repressing both PNC genes were generated using ethanol-inducible RNA interference. A detailed analysis of these plants showed that an impaired peroxisomal ATP import inhibits fatty acid breakdown during early seedling growth and other β-oxidation reactions, such as auxin biosynthesis. We show conclusively that PNC1 and PNC2 are essential for supplying peroxisomes with ATP, indicating that no other ATP generating systems exist inside plant peroxisomes.The β-oxidation of fatty acids, a process that exclusively occurs within peroxisomes in plants and yeast, plays an important role in storage oil mobilization to support seedling establishment of oilseed plants, such as Arabidopsis thaliana (Graham and Eastmond, 2002; Baker et al., 2006; Graham, 2008). Upon germination, fatty acids are released from storage oil triacylglycerol (TAG) by lipolysis, degraded via β-oxidation in specialized peroxisomes, termed glyoxysomes, and subsequently converted to sucrose, which drives growth and development until seedlings become photoautotrophic (Graham and Eastmond, 2002; Baker et al., 2006; Graham, 2008). Before the fatty acids can enter β-oxidation, they are imported into peroxisomes by a peroxisomal ATP binding cassette (ABC) transporter, variously known as CTS (COMATOSE), At PXA1 (Arabidopsis peroxisomal ABC transporter), or PED3 (peroxisomal defective 3) and hereafter referred to as CTS (Zolman et al., 2001; Footitt et al., 2002; Hayashi et al., 2002). Subsequently, the imported fatty acids are activated by esterification to CoA. This ATP-dependent reaction within peroxisomes is catalyzed by long-chain acyl-CoA synthetases 6 and 7 (LACS6 and LACS7, respectively), which are named according to their substrate specificity for long-chain fatty acids, which are significant components of seed storage oil in Arabidopsis (Fulda et al., 2002, 2004).In Saccharomyces cerevisiae, two mechanisms exist for import and activation of fatty acids, depending on chain length (Hettema et al., 1996). Long-chain fatty acids (C16 and C18) are converted to acyl-CoA esters in the cytosol prior to transport by the heterodimeric peroxisomal ABC transporter, Pxa1p/Pxa2 (Hettema et al., 1996). By contrast, short- and medium-chain fatty acids (≤C14) that enter the peroxisomes by passive diffusion or by an unknown transport protein are activated within peroxisomes (Hettema et al., 1996). The possibility cannot be excluded, though, that CTS imports the corresponding CoA derivatives, as is the case for the yeast Pxa1p/Pxa2p heterodimer (Hettema et al., 1996; Verleur et al., 1997), implicating a cytosolic activation of the fatty acids, catalyzed by a hitherto unknown enzyme. The actual substrates transported by CTS in Arabidopsis have not yet been experimentally determined (Theodoulou et al., 2006). However, the sucrose-dependent seedling growth phenotype of the lacs6 lacs7 double knockout mutant demonstrated that peroxisomal activation is essential for lipid mobilization to provide energy for early seedling growth (Fulda et al., 2004). The lacs6 lacs7 mutant is impaired in the degradation of fatty acids, leading to growth arrest shortly after germination (Fulda et al., 2004).Besides fatty acid mobilization, β-oxidation is also involved in generation of signaling molecules, such as the phytohormones auxin and fatty acid–derived jasmonic acid (JA) (Zolman et al., 2000; Schaller et al., 2004; Delker et al., 2007). By analogy to fatty acids released from storage oil, the precursors of these signaling molecules require CoA esterification before they can enter β-oxidation (Baker et al., 2006; Goepfert and Poirier, 2007). While the enzymes responsible for ATP-dependent activation of natural auxin (indole butyric acid [IBA]) and proherbicide 2,4-dichlorophenoxybutyric acid (2,4-DB) are currently unknown, several enzymes belonging to the acyl-activating enzyme (AAE) family have been implicated in jasmonate biosynthesis (Schneider et al., 2005; Koo et al., 2006; Kienow et al., 2008). Moreover, several as yet uncharacterized members of the large AAE family carry a putative peroxisome targeting signal (PTS) and thus might be good candidates to activate the additional β-oxidation substrates within peroxisomes (Shockey et al., 2002, 2003).In the case where activation of fatty acids or other substrates takes place within peroxisomes, the question arises as to how these ATP-dependent reactions are supplied with ATP. It is currently unknown whether plant peroxisomes are able to produce ATP by substrate-level phosphorylation or whether they depend on external ATP to supply energy-dependent reactions within peroxisomes. So far, transport proteins that supply plant peroxisomes with energy for fatty acid oxidation have not been characterized. However, in bakers'' yeast, a peroxisomal adenine nucleotide transporter, ANT1, that is required for the ATP-dependent activation of medium-chain fatty acids inside peroxisomes has been characterized (Palmieri et al., 2001).ATP transport proteins play an important role in the distribution of the primary agent coupling endergonic and exergonic reactions in every cellular compartment (Winkler and Neuhaus, 1999). In Arabidopsis and other plants, various adenine nucleotide carriers have been identified at the molecular level. The mitochondrial ADP/ATP carrier mediates the export of ATP that is synthesized in the mitochondrion to provide energy for cellular metabolism (Heimpel et al., 2001; Haferkamp et al., 2002). The plastidial ATP/ADP transporter (nucleotide transporter) is involved in ATP uptake by both chloroplasts and heterotrophic plastids, to enable the nocturnal ATP supply required for chlorophyll biosynthesis (Reiser et al., 2004; Reinhold et al., 2007), as well as by heterotrophic plastids to drive starch biosynthesis (Batz et al., 1992; Tjaden et al., 1998). Yet another ATP/ADP antiporter located in the endoplasmic reticulum (ER) membrane provides energy by importing ATP into the ER for the accumulation of ER-related storage lipids and proteins (Leroch et al., 2008).In this study, we identified two novel peroxisomal adenine nucleotide carrier proteins (PNC1 and PNC2) from Arabidopsis. Colocalization studies demonstrated that these proteins are targeted to peroxisomes. Yeast complementation and in vitro ATP uptake assays showed that both PNC1 and PNC2 catalyze the counterexchange of ATP with AMP. Using an inducible RNA interference (RNAi) repression strategy, we further established several transgenic Arabidopsis lines with reduced expression levels of both PNC1 and PNC2. Our results showed that import of ATP into peroxisomes that is catalyzed by PNC1 and PNC2 is essential for activation of fatty acids during seedling germination and plays a role in other β-oxidation reactions in peroxisomes, such as auxin metabolism. Analysis of PNC1 and PNC2 repression lines further indicates that no other ATP generating systems exist inside plant peroxisomes and that ATP import is the only way to supply the peroxisomal matrix with ATP.     

Thylakoid Membrane Reduction Affects the Photosystem Stoichiometry in the Cyanobacterium Synechocystis sp. PCC 6803

Biogenesis of thylakoid membranes in both chloroplasts and cyanobacteria is largely not understood today. The vesicle-inducing protein in plastids 1 (Vipp1) has been suggested to be essential for thylakoid membrane formation in Arabidopsis (Arabidopsis thaliana), as well as in the cyanobacterium Synechocystis sp. PCC 6803, although its exact physiological function remains elusive so far. Here, we report that, upon depletion of Vipp1 in Synechocystis cells, the number of thylakoid layers in individual Synechocystis cells decreased, and that, in particular, the content of photosystem I (PSI) complexes was highly diminished in thylakoids. Furthermore, separation of native photosynthetic complexes indicated that PSI trimers are destabilized and the monomeric species is enriched. Therefore, depletion of thylakoid membranes specifically affects biogenesis and/or stabilization of PSI in cyanobacteria.In chloroplasts and cyanobacteria the energy transfer between PSI and PSII is regulated in a light-dependent manner (for a recent review, see Kramer et al., 2004). The two photosystems are connected by the cytochrome b₆f complex, and electron transfer from PSII via the cytochrome b₆f complex to PSI is believed to be regulated by the redox state of the plastoquinol pool potentially also involving the cytochrome b₆f complex (Fujita et al., 1987; Murakami and Fujita, 1993; Schneider et al., 2001, 2004; Pfannschmidt, 2003; Volkmer et al., 2007). Transfer of light energy to the two photosystems is mediated by light-harvesting complexes, and in cyanobacteria light is harvested by the soluble extramembranous phycobilisomes. The efficient energy transfer to PSI and PSII has to be balanced to synchronize the function of the two photosystems. In response to changing light intensities and qualities, energy coupling between the phycobilisomes and the photosystems changes, which allows a rapid adjustment of light absorbance by the individual photosystems. Furthermore, besides this short-term adaptation mechanism, it has been shown in many studies that on a longer term in cyanobacteria the ratio of the two photosystems changes depending on the light conditions (Manodori and Melis, 1986; Murakami and Fujita, 1993; Murakami et al., 1997). Upon shifting cyanobacterial cells from low-light to high-light growth conditions, the PSI-to-PSII ratio decreases due to selective suppression of the amount of functional PSI. In recent years, some genes have already been identified that are involved in this regulation of the photosystem stoichiometry (Hihara et al., 1998; Sonoike et al., 2001; Fujimori et al., 2005; Ozaki et al., 2007).Whereas in chloroplasts of higher plants and green algae the amounts of the two photosystems change in response to changing light conditions (Melis, 1984; Chow et al., 1990; Smith et al., 1990; Kim et al., 1993), it has already been noted a long time ago that the chloroplast ultrastructure also adapts to high-light and low-light conditions (Melis, 1984). Chloroplasts of plants grown under low light or far-red light have more thylakoid membranes than chloroplasts of plants grown under high light or blue light (Anderson et al., 1973; Lichtenthaler et al., 1981; Melis and Harvey, 1981). There appears to be a direct correlation between the chlorophyll content and the amount of thylakoids per chloroplast because light harvesting is increased by enhanced chlorophyll and thylakoid membrane content per chloroplast. Thus, chloroplasts adapt to high light both by a reduction of thylakoid membranes and by a decrease in the PSI-to-PSII ratio.Thylakoid membranes are exclusive features of both cyanobacteria and chloroplasts, and it still remains mysterious how formation of thylakoid membranes is organized. Many cellular processes, like lipid biosynthesis, membrane formation, protein synthesis in the cytoplasm and/or at a membrane, protein transport, protein translocation, and protein folding have to be organized and aligned for formation of internal thylakoid membranes. The recent observation that deletion of the vipp1 gene in Arabidopsis (Arabidopsis thaliana) results in complete loss of thylakoid membranes has indicated that Vipp1 is involved in biogenesis of thylakoid membranes. Further analysis has suggested that Vipp1 could be involved in vesicle trafficking between the inner envelope and the thylakoid membrane of chloroplasts (Kroll et al., 2001). Because of this, the protein was named Vipp1, for vesicle-inducing protein in plastids 1. Depletion of Vipp1 strongly affected the ability of cyanobacterial cells to form proper thylakoid membranes (Westphal et al., 2001) and, consequently, also in cyanobacteria Vipp1 appears to be involved in formation of thylakoid membranes. A Vipp1 depletion strain of Arabidopsis is deficient in photosynthesis, although the defect could not be assigned to a deficiency of a single photosynthetic complex, but appeared to be caused by dysfunction of the entire photosynthetic electron transfer chain (Kroll et al., 2001). Therefore, depletion of Vipp1 in Arabidopsis seems to affect thylakoid membrane formation rather than the assembly of thylakoid membrane protein complexes (Aseeva et al., 2007). However, for cyanobacteria, it is not clear yet how diminishing the amount of thylakoid membrane layers would affect the amount and stoichiometry of the two photosystems.Here, we present the generation and characterization of a Vipp1 depletion strain of the cyanobacterium Synechocystis sp. PCC 6803. Upon depletion of Vipp1, a decrease in thylakoid membrane pairs in the generated mutant strain and, furthermore, a significant decrease in active PSI centers was observed. Moreover, trimerization of PSI also appeared to be impaired in the mutant strain. These results suggest that thylakoid membrane perturbations caused by the Vipp1 depletion directly affects PSI assembly and stability in cyanobacterial thylakoid membranes.     

Inflammation in Response to n3 Fatty Acids in a Porcine Obesity Model

Fatty acids have distinct cellular effects related to inflammation and insulin sensitivity. Dietary saturated fat activates toll-like receptor 4, which in turn can lead to chronic inflammation, insulin resistance, and adipose tissue macrophage infiltration. Conversely, n3 fatty acids are generally antiinflammatory and promote insulin sensitivity, in part via peroxisome proliferator-activated receptor γ. Ossabaw swine are a useful biomedical model of obesity. We fed Ossabaw pigs either a low-fat control diet or a diet containing high-fat palm oil with or without additional n3 fatty acids for 30 wk to investigate the effect of saturated fats and n3 fatty acids on obesity-linked inflammatory markers. The diet did not influence the inflammatory markers C-reactive protein, TNFα, IL6, or IL12. In addition, n3 fatty acids attenuated the increase in inflammatory adipose tissue CD16^–CD14⁺ macrophages induced by high palm oil. High-fat diets with and without n3 fatty acids both induced hyperglycemia without hyperinsulinemia. The high-fat only group but not the high-fat group with n3 fatty acids showed reduced insulin sensitivity in response to insulin challenge. This effect was not mediated by decreased phosphorylation of protein kinase B. Therefore, in obese Ossabaw swine, n3 fatty acids partially attenuate insulin resistance but only marginally change inflammatory status and macrophage phenotype in adipose tissue.Abbreviations: AMPKα, AMP-activated protein kinase α, CRP, C-reactive protein, DHA, docosahexanoic acid, EPA, eicosapentanoic acid, HFP, high-fat palm-oil diet, HFPn3, high-fat palm-oil diet supplemented with n3 fatty acids, HOMA-IR, homeostasis model of assessment–, insulin resistance, LFC, low-fat control diet, PKB, protein kinase B, PUFA, polyunsaturated fatty acidsObesity is accompanied by chronic inflammation in adipose tissue; increased circulating concentrations of TNFα, IL6, and C-reactive protein (CRP); and decreased concentrations of adiponectin.² This chronic inflammation links obesity and the development of insulin resistance.³⁹ Dietary saturated fatty acids promote obesity in part through the induction of inflammation via activation of toll-like receptor 4 (the innate immune receptor for LPS).²⁸ The absence of functional tlr4 in mice reduces circulating proinflammatory cytokine concentrations and decreases macrophage infiltration into adipose tissue during high-fat diet-induced obesity.^8,28,32 Furthermore, in 3T3 L1 mouse adipocytes, palmitate activates NFκB, protein kinase C, and mitogen-activated protein kinase, all of which increase the production of inflammatory cytokines.¹For people who consume a diet high in saturated fat, a major determinant of health is the ratio of omega-6 to omega-3 fatty acids (that is, n6:n3) that is consumed.^5,6 Unlike saturated fatty acids, the n3 polyunsaturated fatty acids (PUFA) eicosapentanoic acid (EPA) and docosahexanoic acid (DHA) exert predominantly antiinflammatory effects, as is evident in that DHA antagonizes NFκB activation by palmitate in 3T3 L1 adipocytes.¹ In mice, EPA prevents or reverses hyperinsulinemia, hyperglycemia, and increased circulating monocyte chemotatic protein 1¹⁶ and decreases infiltration of adipose tissue with macrophages.³⁰ Moreover, n3 PUFA alleviate the decline in serum adiponectin that is associated with obesity,^12,15,30 and EPA decreases serum CRP in diabetic patients.²⁶Physiologic differences between rodents and humans underscore the need for comparative models in biomedical research, and the pig is emerging rapidly as a model for studies of energy metabolism and obesity. Like humans, pigs are natural omnivores, rely on apolipoprotein B100 to shuttle cholesterol in the LDL fraction, and have minimal brown fat retention postnatally. Furthermore, adipose depots in pigs are of sufficient size that multiple assays can be done on adipocytes or stromal vascular cells without pooling across depots or animals. Although Ossabaw swine have been used as models for metabolic syndrome, cardiovascular disease, coronary artery disease, and steatohepatisis,^11,20,24 little is known about adipose inflammation in these animals. Consequently, we sought to characterize obesity-linked inflammatory markers in the adipose tissue of this novel model and to test the hypothesis that adding n3 PUFA to a diet high in saturated fat attenuates chronic inflammation, protects against diet-induced insulin resistance, and alters phenotypic changes in adipose tissue macrophages.     

PLASTID MOVEMENT IMPAIRED1 and PLASTID MOVEMENT IMPAIRED1-RELATED1 Mediate Photorelocation Movements of Both Chloroplasts and Nuclei

Organelle movement and positioning play important roles in fundamental cellular activities and adaptive responses to environmental stress in plants. To optimize photosynthetic light utilization, chloroplasts move toward weak blue light (the accumulation response) and escape from strong blue light (the avoidance response). Nuclei also move in response to strong blue light by utilizing the light-induced movement of attached plastids in leaf cells. Blue light receptor phototropins and several factors for chloroplast photorelocation movement have been identified through molecular genetic analysis of Arabidopsis (Arabidopsis thaliana). PLASTID MOVEMENT IMPAIRED1 (PMI1) is a plant-specific C2-domain protein that is required for efficient chloroplast photorelocation movement. There are two PLASTID MOVEMENT IMPAIRED1-RELATED (PMIR) genes, PMIR1 and PMIR2, in the Arabidopsis genome. However, the mechanism in which PMI1 regulates chloroplast and nuclear photorelocation movements and the involvement of PMIR1 and PMIR2 in these organelle movements remained unknown. Here, we analyzed chloroplast and nuclear photorelocation movements in mutant lines of PMI1, PMIR1, and PMIR2. In mesophyll cells, the pmi1 single mutant showed severe defects in both chloroplast and nuclear photorelocation movements resulting from the impaired regulation of chloroplast-actin filaments. In pavement cells, pmi1 mutant plants were partially defective in both plastid and nuclear photorelocation movements, but pmi1pmir1 and pmi1pmir1pmir2 mutant lines lacked the blue light-induced movement responses of plastids and nuclei completely. These results indicated that PMI1 is essential for chloroplast and nuclear photorelocation movements in mesophyll cells and that both PMI1 and PMIR1 are indispensable for photorelocation movements of plastids and thus, nuclei in pavement cells.In plants, organelles move within the cell and become appropriately positioned to accomplish their functions and adapt to the environment (for review, see Wada and Suetsugu, 2004). Light-induced chloroplast movement (chloroplast photorelocation movement) is one of the best characterized organelle movements in plants (Suetsugu and Wada, 2012). Under weak light conditions, chloroplasts move toward light to capture light efficiently (the accumulation response; Zurzycki, 1955). Under strong light conditions, chloroplasts escape from light to avoid photodamage (the avoidance response; Kasahara et al., 2002; Sztatelman et al., 2010; Davis and Hangarter, 2012; Cazzaniga et al., 2013). In most green plant species, these responses are induced primarily by the blue light receptor phototropin (phot) in response to a range of wavelengths from UVA to blue light (approximately 320–500 nm; for review, see Suetsugu and Wada, 2012; Wada and Suetsugu, 2013; Kong and Wada, 2014). Phot-mediated chloroplast movement has been shown in land plants, such as Arabidopsis (Arabidopsis thaliana; Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001), the fern Adiantum capillus-veneris (Kagawa et al., 2004), the moss Physcomitrella patens (Kasahara et al., 2004), and the liverwort Marchantia polymorpha (Komatsu et al., 2014). Two phots in Arabidopsis, phot1 and phot2, redundantly mediate the accumulation response (Sakai et al., 2001), whereas phot2 primarily regulates the avoidance response (Jarillo et al., 2001; Kagawa et al., 2001; Luesse et al., 2010). M. polymorpha has only one phot that mediates both the accumulation and avoidance responses (Komatsu et al., 2014), although two or more phots mediate chloroplast photorelocation movement in A. capillus-veneris (Kagawa et al., 2004) and P. patens (Kasahara et al., 2004). Thus, duplication and functional diversification of PHOT genes have occurred during land plant evolution, and plants have gained a sophisticated light sensing system for chloroplast photorelocation movement.In general, movements of plant organelles, including chloroplasts, are dependent on actin filaments (for review, see Wada and Suetsugu, 2004). Most organelles common in eukaryotes, such as mitochondria, peroxisomes, and Golgi bodies, use the myosin motor for their movements, but there is no clear evidence that chloroplast movement is myosin dependent (for review, see Suetsugu et al., 2010a). Land plants have innovated a novel actin-based motility system that is specialized for chloroplast movement as well as a photoreceptor system (for review, see Suetsugu et al., 2010a; Wada and Suetsugu, 2013; Kong and Wada, 2014). Chloroplast-actin (cp-actin) filaments, which were first found in Arabidopsis, are short actin filaments specifically localized around the chloroplast periphery at the interface between the chloroplast and the plasma membrane (Kadota et al., 2009). Strong blue light induces the rapid disappearance of cp-actin filaments and then, their subsequent reappearance preferentially at the front region of the moving chloroplasts. This asymmetric distribution of cp-actin filaments is essential for directional chloroplast movement (Kadota et al., 2009; Kong et al., 2013a). The greater the difference in the amount of cp-actin filaments between the front and rear regions of chloroplasts becomes, the faster the chloroplasts move, in which the magnitude of the difference is determined by fluence rate (Kagawa and Wada, 2004; Kadota et al., 2009; Kong et al., 2013a). Strong blue light-induced disappearance of cp-actin filaments is regulated in a phot2-dependent manner before the intensive polymerization of cp-actin filaments at the front region occurs (Kadota et al., 2009; Ichikawa et al., 2011; Kong et al., 2013a). This phot2-dependent response contributes to the greater difference in the amount of cp-actin filaments between the front and rear regions of chloroplasts. Similar behavior of cp-actin filaments has also been observed in A. capillus-veneris (Tsuboi and Wada, 2012) and P. patens (Yamashita et al., 2011).Like chloroplasts, nuclei also show light-mediated movement and positioning (nuclear photorelocation movement) in land plants (for review, see Higa et al., 2014b). In gametophytic cells of A. capillus-veneris, weak light induced the accumulation responses of both chloroplasts and nuclei, whereas strong light induced avoidance responses (Kagawa and Wada, 1993, 1995; Tsuboi et al., 2007). However, in mesophyll cells of Arabidopsis, strong blue light induced both chloroplast and nuclear avoidance responses, but weak blue light induced only the chloroplast accumulation response (Iwabuchi et al., 2007, 2010; Higa et al., 2014a). In Arabidopsis pavement cells, small numbers of tiny plastids were found and showed autofluorescence under the confocal laser-scanning microscopy (Iwabuchi et al., 2010; Higa et al., 2014a). Hereafter, the plastid in the pavement cells is called the pavement cell plastid. Strong blue light-induced avoidance responses of pavement cell plastids and nuclei were induced in a phot2-dependent manner, but the accumulation response was not detected for either organelle (Iwabuchi et al., 2007, 2010; Higa et al., 2014a). In both Arabidopsis and A. capillus-veneris, phots mediate nuclear photorelocation movement, and phot2 mediates the nuclear avoidance response (Iwabuchi et al., 2007, 2010; Tsuboi et al., 2007). The nuclear avoidance response is dependent on actin filaments in both mesophyll and pavement cells of Arabidopsis (Iwabuchi et al., 2010). Recently, it was shown that the nuclear avoidance response relies on cp-actin-dependent movement of pavement cell plastids, where nuclei are associated with pavement cell plastids of Arabidopsis (Higa et al., 2014a). In mesophyll cells, nuclear avoidance response is likely dependent on cp-actin filament-mediated chloroplast movement, because the mutants deficient in chloroplast movement were also defective in nuclear avoidance response (Higa et al., 2014a). Thus, phots mediate both chloroplast (and pavement cell plastid) and nuclear photorelocation movement by regulating cp-actin filaments.Molecular genetic analyses of Arabidopsis mutants deficient in chloroplast photorelocation movement have identified many molecular factors involved in signal transduction and/or motility systems as well as those involved in the photoreceptor system for chloroplast photorelocation movement (and thus, nuclear photorelocation movement; for review, see Suetsugu and Wada, 2012; Wada and Suetsugu, 2013; Kong and Wada, 2014). CHLOROPLAST UNUSUAL POSITIONING1 (CHUP1; Oikawa et al., 2003) and KINESIN-LIKE PROTEIN FOR ACTIN-BASED CHLOROPLAST MOVEMENT (KAC; Suetsugu et al., 2010b) are key factors for generating and/or maintaining cp-actin filaments. Both proteins are highly conserved in land plants and essential for the movement and attachment of chloroplasts to the plasma membrane in Arabidopsis (Oikawa et al., 2003, 2008; Suetsugu et al., 2010b), A. capillus-veneris (Suetsugu et al., 2012), and P. patens (Suetsugu et al., 2012; Usami et al., 2012). CHUP1 is localized on the chloroplast outer membrane and binds to globular and filamentous actins and profilin in vitro (Oikawa et al., 2003, 2008; Schmidt von Braun and Schleiff, 2008). Although KAC is a kinesin-like protein, it lacks microtubule-dependent motor activity but has filamentous actin binding activity (Suetsugu et al., 2010b). An actin-bundling protein THRUMIN1 (THRUM1) is required for efficient chloroplast photorelocation movement (Whippo et al., 2011) and interacts with cp-actin filaments (Kong et al., 2013a). chup1 and kac mutant plants were shown to lack detectable cp-actin filaments (Kadota et al., 2009; Suetsugu et al., 2010b; Ichikawa et al., 2011; Kong et al., 2013a). Similarly, cp-actin filaments were rarely detected in thrum1 mutant plants (Kong et al., 2013a), indicating that THRUM1 also plays an important role in maintaining cp-actin filaments.Other proteins J-DOMAIN PROTEIN REQUIRED FOR CHLOROPLAST ACCUMULATION RESPONSE1 (JAC1; Suetsugu et al., 2005), WEAK CHLOROPLAST MOVEMENT UNDER BLUE LIGHT1 (WEB1; Kodama et al., 2010), and PLASTID MOVEMENT IMPAIRED2 (PMI2; Luesse et al., 2006; Kodama et al., 2010) are involved in the light regulation of cp-actin filaments and chloroplast photorelocation movement. JAC1 is an auxilin-like J-domain protein that mediates the chloroplast accumulation response through its J-domain function (Suetsugu et al., 2005; Takano et al., 2010). WEB1 and PMI2 are coiled-coil proteins that interact with each other (Kodama et al., 2010). Although web1 and pmi2 were partially defective in the avoidance response, the jac1 mutation completely suppressed the phenotype of web1 and pmi2, suggesting that the WEB1/PMI2 complex suppresses JAC1 function (i.e. the accumulation response) under strong light conditions (Kodama et al., 2010). Both web1 and pmi2 showed impaired disappearance of cp-actin filaments in response to strong blue light (Kodama et al., 2010). However, the exact molecular functions of these proteins are unknown.In this study, we characterized mutant plants deficient in the PMI1 gene and two homologous genes PLASTID MOVEMENT IMPAIRED1-RELATED1 (PMIR1) and PMIR2. PMI1 was identified through molecular genetic analyses of pmi1 mutants that showed severe defects in chloroplast accumulation and avoidance responses (DeBlasio et al., 2005). PMI1 is a plant-specific C2-domain protein (DeBlasio et al., 2005; Zhang and Aravind, 2010), but its roles and those of PMIRs in cp-actin-mediated chloroplast and nuclear photorelocation movements remained unclear. Thus, we analyzed chloroplast and nuclear photorelocation movements in the single, double, and triple mutants of pmi1, pmir1, and pmir2.     

New-Onset Diabetes Mellitus After Transplantation in a Cynomolgus Macaque (Macaca fasicularis)

A 5.5-y-old intact male cynomolgus macaque (Macaca fasicularis) presented with inappetence and weight loss 57 d after heterotopic heart and thymus transplantation while receiving an immunosuppressant regimen consisting of tacrolimus, mycophenolate mofetil, and methylprednisolone to prevent graft rejection. A serum chemistry panel, a glycated hemoglobin test, and urinalysis performed at presentation revealed elevated blood glucose and glycated hemoglobin (HbA1c) levels (727 mg/dL and 10.1%, respectively), glucosuria, and ketonuria. Diabetes mellitus was diagnosed, and insulin therapy was initiated immediately. The macaque was weaned off the immunosuppressive therapy as his clinical condition improved and stabilized. Approximately 74 d after discontinuation of the immunosuppressants, the blood glucose normalized, and the insulin therapy was stopped. The animal''s blood glucose and HbA1c values have remained within normal limits since this time. We suspect that our macaque experienced new-onset diabetes mellitus after transplantation, a condition that is commonly observed in human transplant patients but not well described in NHP. To our knowledge, this report represents the first documented case of new-onset diabetes mellitus after transplantation in a cynomolgus macaque.Abbreviations: NODAT, new-onset diabetes mellitus after transplantationNew-onset diabetes mellitus after transplantation (NODAT, formerly known as posttransplantation diabetes mellitus) is an important consequence of solid-organ transplantation in humans.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} A variety of risk factors have been identified including increased age, sex (male prevalence), elevated pretransplant fasting plasma glucose levels, and immunosuppressive therapy.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} The relationship between calcineurin inhibitors, such as tacrolimus and cyclosporin, and the development of NODAT is widely recognized in human medicine.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} Cynomolgus macaques (Macaca fasicularis) are a commonly used NHP model in organ transplantation research. Cases of natural and induced diabetes of cynomolgus monkeys have been described in the literature;^14,43,45 however, NODAT in a macaque model of solid-organ transplantation has not been reported previously to our knowledge.     

Chloroplast DNA Replication Is Regulated by the Redox State Independently of Chloroplast Division in Chlamydomonas reinhardtii

Chloroplasts arose from a cyanobacterial endosymbiont and multiply by division. In algal cells, chloroplast division is regulated by the cell cycle so as to occur only once, in the S phase. Chloroplasts possess multiple copies of their own genome that must be replicated during chloroplast proliferation. In order to examine how chloroplast DNA replication is regulated in the green alga Chlamydomonas reinhardtii, we first asked whether it is regulated by the cell cycle, as is the case for chloroplast division. Chloroplast DNA is replicated in the light and not the dark phase, independent of the cell cycle or the timing of chloroplast division in photoautotrophic culture. Inhibition of photosynthetic electron transfer blocked chloroplast DNA replication. However, chloroplast DNA was replicated when the cells were grown heterotrophically in the dark, raising the possibility that chloroplast DNA replication is coupled with the reducing power supplied by photosynthesis or the uptake of acetate. When dimethylthiourea, a reactive oxygen species scavenger, was added to the photoautotrophic culture, chloroplast DNA was replicated even in the dark. In contrast, when methylviologen, a reactive oxygen species inducer, was added, chloroplast DNA was not replicated in the light. Moreover, the chloroplast DNA replication activity in both the isolated chloroplasts and nucleoids was increased by dithiothreitol, while it was repressed by diamide, a specific thiol-oxidizing reagent. These results suggest that chloroplast DNA replication is regulated by the redox state that is sensed by the nucleoids and that the disulfide bonds in nucleoid-associated proteins are involved in this regulatory activity.Chloroplasts are semiautonomous organelles that possess their own genome, which is complexed with proteins to form nucleoids and also certain machinery needed for protein synthesis, as is the case in prokaryotes. It is generally accepted that chloroplasts arose from a bacterial endosymbiont closely related to the currently extant cyanobacteria (Archibald, 2009; Keeling, 2010). In a manner reminiscent of their free-living ancestor, chloroplasts proliferate by the division of preexisting organelles that are coupled to the duplication and segregation of the nucleoids (Kuroiwa, 1991) and have retained the bulk of their bacterial biochemistry. However, chloroplasts have subsequently been substantially remodeled by the host cell so as to function as complementary organelles within the eukaryotic host cell (Rodríguez-Ezpeleta and Philippe, 2006; Archibald, 2009; Keeling, 2010). For example, most of the genes that were once in the original endosymbiont genome have been either lost or transferred into the host nuclear genome. As a result, the size of the chloroplast genome has been reduced to less than one-tenth that of the free-living cyanobacterial genome. Thus, the bulk of the chloroplast proteome consists of nucleus-encoded proteins that are translated on cytoplasmic ribosomes and translocated into chloroplasts. In addition, chloroplast division ultimately came to be a process tightly regulated by the host cell, which ensured permanent inheritance of the chloroplasts during the course of cell division and from generation to generation (Rodríguez-Ezpeleta and Philippe, 2006; Archibald, 2009; Keeling, 2010).Chloroplast division is performed by constriction of the ring structures at the division site, encompassing both the inside and the outside of the two envelopes (Yang et al., 2008; Maple and Møller, 2010; Miyagishima, 2011; Pyke, 2013). One part of the division machinery is derived from the cyanobacterial cytokinetic machinery that is based on the FtsZ protein. In contrast, other parts of the division machinery involve proteins specific to eukaryotes, including one member of the dynamin family. The majority of algae (both unicellular and multicellular), which diverged early within the Plantae, have just one or at most only a few chloroplasts per cell. In algae, the chloroplast divides once per cell cycle before the host cell completes cytokinesis (Suzuki et al., 1994; Miyagishima et al., 2012). In contrast, land plants and certain algal species contain dozens of chloroplasts per cell that divide nonsynchronously, even within the same cell (Boffey and Lloyd, 1988). Because land plants evolved from algae, there is likely to have been a linkage between the cell cycle and chloroplast division in their algal ancestor that was subsequently lost during land plant evolution. Our recent study showed that the timing of chloroplast division in algae is restricted to the S phase by S phase-specific formation of the chloroplast division machinery, which is based on the cell cycle-regulated expression of the components of the chloroplast division machinery (Miyagishima et al., 2012).Because chloroplasts possess their own genome, chloroplast DNA must be duplicated so that each daughter chloroplast inherits the required DNA after division. However, it is still unclear how the replication of chloroplast DNA is regulated and whether the replication is coupled with the timing of chloroplast division, even though certain studies have addressed this issue, as described below.Bacteria such as Escherichia coli and Bacillus subtilis possess a single circular chromosome. In these bacteria, the process of DNA replication is tightly coupled with cell division (Boye et al., 2000; Zakrzewska-Czerwińska et al., 2007), in which the initiation of replication is regulated such that it occurs only once per cell division cycle (Boye et al., 2000). In contrast, cyanobacteria contain multiple copies of their DNA (e.g. three to five copies in Synechococcus elongatus PCC 7942; Mann and Carr, 1974; Griese et al., 2011). In some obligate photoautotrophic cyanobacterial species, replication is initiated only when light is available (Binder and Chisholm, 1990; Mori et al., 1996; Watanabe et al., 2012). Replication is initiated asynchronously among the multiple copies of the DNA. Although the regulation of the initiation of DNA replication is less stringent than that in E. coli and B. subtilis, as described above, a recent study using S. elongatus PCC 7942 showed that this replication peaks prior to cell division, as in other bacteria.Chloroplasts also contain multiple copies of DNA (approximately 1,000 copies; Boffey and Leech, 1982; Miyamura et al., 1986; Baumgartner et al., 1989; Oldenburg and Bendich, 2004; Oldenburg et al., 2006; Shaver et al., 2008). In algae, chloroplast DNA is replicated in a manner that keeps pace with chloroplast and cell division in order to maintain the proper DNA content per chloroplast (i.e. per cell). In contrast, in land plants, the copy number of DNA in each chloroplast (plastid) changes during the course of development and differentiation, although contradictory results were reported about leaf development (Lamppa and Bendich, 1979; Boffey and Leech, 1982; Hashimoto and Possingham, 1989; Kuroiwa, 1991; Rowan and Bendich, 2009; Matsushima et al., 2011). Previous studies that synchronized the algal cell cycle by means of a 24-h light/dark cycle showed that chloroplast DNA is replicated only during the G1 phase, after which it is separated into daughter chloroplasts during the S phase by chloroplast division, implying that chloroplast DNA replication and division are temporally separated (Chiang and Sueoka, 1967; Grant et al., 1978; Suzuki et al., 1994). However, under these experimental conditions, G1 cells grow and the chloroplast DNA level increases during the light period. Cells enter into the S phase, chloroplast DNA replication ceases, and the chloroplasts divide at the beginning of the dark period. Thus, it is still unclear whether chloroplast DNA replication is directly controlled by the cell cycle, as is the case in chloroplast division, or chloroplast DNA replication occurs merely when light energy is available.We addressed this issue using a synchronous culture as well as a heterotrophic culture of the mixotrophic green alga Chlamydomonas reinhardtii. The results show that chloroplast DNA replication occurs independently of either the cell cycle or the timing of chloroplast division. Instead, it is shown that chloroplast DNA replication occurs when light is available in photoautotrophic culture and even under darkness in heterotrophic culture. Further experimental results suggest that chloroplast DNA replication is regulated by the redox state in the cell, which is sensed by the chloroplast nucleoids.     

Abscisic Acid Deficiency Causes Changes in Cuticle Permeability and Pectin Composition That Influence Tomato Resistance to Botrytis cinerea

A mutant of tomato (Solanum lycopersicum) with reduced abscisic acid (ABA) production (sitiens) exhibits increased resistance to the necrotrophic fungus Botrytis cinerea. This resistance is correlated with a rapid and strong hydrogen peroxide-driven cell wall fortification response in epidermis cells that is absent in tomato with normal ABA production. Moreover, basal expression of defense genes is higher in the mutant compared with the wild-type tomato. Given the importance of this fast response in sitiens resistance, we investigated cell wall and cuticle properties of the mutant at the chemical, histological, and ultrastructural levels. We demonstrate that ABA deficiency in the mutant leads to increased cuticle permeability, which is positively correlated with disease resistance. Furthermore, perturbation of ABA levels affects pectin composition. sitiens plants have a relatively higher degree of pectin methylesterification and release different oligosaccharides upon inoculation with B. cinerea. These results show that endogenous plant ABA levels affect the composition of the tomato cuticle and cell wall and demonstrate the importance of cuticle and cell wall chemistry in shaping the outcome of this plant-fungus interaction.Plant defense against pathogens often involves the induction of mechanisms after pathogen recognition, including defense signaling, cell wall strengthening, and localized cell death, but plants also have preformed chemical and structural defense barriers. Fungal pathogens that penetrate the plant tissue directly through the outer surface, rather than via natural plant openings or wounds, must pass through the plant cuticle and epidermal cell wall. Penetration of the host surface happens either by physical means (i.e. by a highly localized pressure in the appressorium) or by chemical means (i.e. by the release of hydrolyzing enzymes). Necrotrophic plant pathogens like Botrytis cinerea typically use the latter strategy. During penetration, they produce cutinases and pectinolytic enzymes such as pectin methylesterases, endopolygalacturonases, and exopolygalacturonases (van Kan, 2006).The cuticle is a hydrophobic barrier that covers the aerial surfaces of the plant. It is mainly composed of cutin, a polyester matrix, and soluble waxes, a complex mixture of hydrophobic material containing very-long-chain fatty acids and their derivatives, embedded into and deposited onto the cutin matrix. It plays an important role in organ development and protection against water loss (Yephremov et al., 1999; Sieber et al., 2000; Kurata et al., 2003; Jung et al., 2006). The cuticle is generally considered as a mere passive physical barrier against pathogen invasion, but it has also been recognized as a potential source of signaling and elicitor molecules (Jenks et al., 1994; Reina-Pinto and Yephremov, 2009). Plant cutin monomers trigger cutinase secretion in pathogenic fungi (Woloshuk and Kolattukudy, 1986), and cutin and wax components initiate appressorium formation and penetration in appressorium-forming pathogens (Kolattukudy et al., 1995; Francis et al., 1996; Gilbert et al., 1996; Fauth et al., 1998; Dickman et al., 2003). In plants, cutin monomers induce pathogenesis-related gene expression and elicit hydrogen peroxide (H₂O₂) synthesis (Fauth et al., 1998; Kim et al., 2008; Park et al., 2008). Transgenic tomato (Solanum lycopersicum) plants expressing the yeast Δ-9 desaturase gene had high levels of cutin monomers that inhibited powdery mildew (Erysiphe polygoni) spore germination, leading to enhanced resistance (Wang et al., 2000). Arabidopsis (Arabidopsis thaliana) plants expressing a fungal cutinase or mutants with a defective cuticle, such as long-chain acyl-CoA synthetase2 and bodyguard, are generally more susceptible to bacteria and equally susceptible to biotrophic fungi but are surprisingly resistant to B. cinerea (Bessire et al., 2007; Chassot et al., 2007; Tang et al., 2007). It has been postulated that a defective or thin cuticle encourages these plants to constitutively express defense-related mechanisms and to secrete antifungal compounds to the plant surface, thereby inhibiting B. cinerea growth (Bessire et al., 2007; Chassot et al., 2007). In addition, cuticle metabolic pathways might directly modulate plant-pathogen interactions by interacting with hormonally regulated defense pathways (Fiebig et al., 2000; Garbay et al., 2007; Mang et al., 2009) or with complex lipid signaling pathways leading to hypersensitive cell death (Raffaele et al., 2008).Once plant pathogens have penetrated the cuticle, they secrete hydrolases that target the plant cell wall (ten Have et al., 1998; Oeser et al., 2002; Vogel et al., 2002; Jakob et al., 2007) that is mainly composed of cellulose, hemicellulose, and pectin (35% of total dry weight). Pectin consists mainly of the polysaccharides homogalacturonan and rhamnogalacturonan I and II. Homogalacturonans are linear chains of α-(1–4)-linked d-GalA residues that can be methylesterified at C-6. Rhamnogalacturonan I and II are more complex, branched polysaccharides. B. cinerea is typically regarded as a pectinolytic pathogen because it possesses an efficient pectinolytic machinery, including a variety of polygalacturonases and pectin methylesterases (PMEs), some of which are important virulence factors (ten Have et al., 1998, 2001; Valette-Collet et al., 2003; Kars et al., 2005). Pectins are a rich source of oligogalacturonides (OGAs), biologically active signaling molecules that can activate plant defense mechanisms (Hahn et al., 1981; Côté and Hahn, 1994; Messiaen and Van Cutsem, 1994; Ridley et al., 2001). The eliciting capacity of the OGAs was shown to depend on their size, which in turn is influenced by the methylesterification pattern of the homogalacturonan fraction (Mathieu et al., 1991; Messiaen and Van Cutsem, 1994). To counteract the activity of fungal pectinases, many plants express polygalacturonase-inhibiting proteins and PME inhibitors, which are localized in the cell wall. The role of these proteins in plant defense against B. cinerea has been extensively demonstrated (Powell et al., 2000; Ferrari et al., 2003; Sicilia et al., 2005; Joubert et al., 2006, 2007; Lionetti et al., 2007). The interaction with the inhibitors not only limits the destructive potential of polygalacturonases but also leads to the accumulation of elicitor-active OGAs (De Lorenzo and Ferrari, 2002). How OGAs are perceived by the plant is still unclear, but in view of the diversity of biological activities and structure requirements, they are thought to be recognized through different proteins, including receptor-like kinases, wall-associated kinases, arabinogalactan proteins, and Pro-rich proteins (Côté and Hahn, 1994; Showalter, 2001; Humphrey et al., 2007).Over the past years, the role of abscisic acid (ABA) in plant-pathogen interactions has gained increased attention. ABA is mostly negatively correlated with resistance against phytopathogens through down-regulation of defense responses orchestrated by salicylic acid, jasmonic acid, and ethylene (Mohr and Cahill, 2001; Audenaert et al., 2002; Mauch-Mani and Mauch, 2005; Asselbergh et al., 2008). In tomato, the ABA-deficient mutant sitiens has an enhanced resistance to B. cinerea (Audenaert et al., 2002) that depends on a timely, localized oxidative burst leading to rapid epidermal cell wall fortification and a faster and higher induction of defense-related gene expression upon infection compared with the wild type (Asselbergh et al., 2007). Moreover, basal defense gene expression is higher in this mutant than in the wild type. As this early response is of vital importance for the resistant reaction of tomato against B. cinerea, we investigated whether alterations in cuticle and/or cell wall, which form the first barrier to the invading pathogen, affect resistance. We demonstrate that the sitiens cuticle is more permeable and that permeability is positively correlated with resistance to B. cinerea. Furthermore, differences in pectin composition and rate of methylesterification occur. Together, these data hint at an unanticipated role for extracellular matrix components in the resistance of tomato against B. cinerea and thus shed new light on the largely unexplored interrelationship between the extracellular matrix and plant-pathogen interactions.