Three Fatty Acyl-Coenzyme A Reductases, BdFAR1, BdFAR2 and BdFAR3, are Involved in Cuticular Wax Primary Alcohol Biosynthesis in Brachypodium distachyon

Three Fatty Acyl-Coenzyme A Reductases, BdFAR1, BdFAR2 and BdFAR3, are Involved in Cuticular Wax... Abstract Plant cuticular wax is a heterogeneous mixture of very long chain fatty acids (VLCFAs) and their derivatives. Primary alcohols are the dominant wax components throughout leaf development of Brachypodium distachyon (Brachypodium). However, the genes involved in primary alcohol biosynthesis have not been investigated and their exact biological function remains unclear in Brachypodium to date. Here, we monitored the leaf wax profile and crystal morphology during Brachypodium leaf morphogenesis, and isolated three Brachypodium fatty acyl-CoA reductase (FAR) genes, named BdFAR1, BdFAR2 and BdFAR3, then analyzed their biochemical activities, substrate specificities, expression patterns, subcellular localization and stress induction. Transgenic expression of BdFAR genes in yeast (Saccharomyces cerevisiae), tomato (Solanum lycopersicum), Arabidopsis (Arabidopsis thaliana) and Brachypodium increased the production of primary alcohols. The three BdFAR genes were preferentially expressed in Brachypodium aerial tissues, consistent with known sites of wax primary alcohol deposition, and localized in the endoplasmic reticulum (ER) in Arabidopsis protoplasts. Finally, expression of the BdFAR genes was induced by drought, cold and ABA treatments, and drought stress significantly increased cuticular wax accumulation in Brachypodium. Taken together, these results indicate that the three BdFAR genes encode active FARs involved in the biosynthesis of Brachypodium wax primary alcohols and respond to abiotic stresses. Introduction The cuticle is a continuous hydrophobic layer that coats most aerial surfaces of terrestrial plants and forms the contact zone between the plant and the environment. The cuticle plays a critical role in limiting non-stomatal water loss (Yeats and Rose 2013) and helps to protect plants against particulate deposits (Barthlott and Neinhuis 1997), UV radiation (Barnes et al. 1996), and bacterial and fungal pathogens (Raffaele et al. 2009). Structurally, the cuticle comprises two major components, cutin and wax. Cutin is a three-dimensional polyester primarily composed of C16 and C18 hydroxy fatty acids, epoxy fatty acids, diacids and glycerol (Beisson et al. 2012), while cuticular wax is a complex mixture commonly including very long chain fatty acids (VLCFAs), primary and secondary alcohols, ketones, aldehydes, alkanes and alkyl esters (Jetter et al. 2007, Samuels et al. 2008). The relative composition of cuticular wax varies greatly among plant species, tissues and different developmental stages of the same organ (Eigenbrode and Espelie 1995, Y. Wang et al. 2015a). The cuticle surfaces of many species and organs are smooth, resulting in a glossy macroscopic appearance, whereas the cuticle surfaces of other species have protruding microscopic wax crystals that give the surface a glaucous appearance (Jeffree 2007). Scanning electron microscopy (SEM) is commonly used to distinguish characteristic wax crystal shapes including platelets, tubules, rodlets and rods (Barthlott et al. 1998), which correlated with certain wax constituents (Jeffree 2007). In diverse plant species and organs, the most important wax compounds are primary alcohols with predominant chain lengths of C26–C30 (Baker 1982). Previous studies have shown that wax alcohols are produced by the acyl reduction pathway in Brassica oleracea L. and involve two separate enzymes with aldehydes as intermediates (Kunst and Samuels 2003). However, subsequent studies have demonstrated that, in Arabidopsis thaliana (Arabidopsis), a single fatty acyl-CoA reductase (FAR) can directly reduce VLC acyl-CoA precursors to primary alcohols (Kunst and Samuels 2003, Samuels et al. 2008). Functional expression of FARs from jojoba (Metz et al. 2000), silkmoth (Bombyx mori) (Moto et al. 2003), mouse (Mus musculus), human (Homo sapiens) (Cheng and Russell 2004), Euglena gracilis (Teerawanichpan and Qiu, 2010) and Marinobacter aquaeolei (Hofvander et al. 2011, Willis et al. 2011, Fillet et al. 2015, W. Wang et al. 2016) in heterologous systems further supported this idea. In recent years, there has been increased interest in FAR proteins that produce primary alcohols. In Arabidopsis, a family of eight putative FAR-like proteins has been identified. AtMS2/FAR2 is involved in the formation of fatty alcohols during pollen exine development (Aarts et al. 1997, Doan et al. 2009, Dobritsa et al. 2009, Chen et al. 2011), while AtCER4/FAR3 catalyzes the synthesis of C24–C30 primary alcohols found in cuticular wax of aerial organs (Rowland et al. 2006). AtFAR1, AtFAR4 and AtFAR5 generate the fatty alcohols found in root, seed coat and wound-induced leaf tissue (Domergue et al. 2010). In wheat, the anther-specific protein TAA1a synthesizes fatty alcohols (Wang et al. 2002). TaFAR1 and TaFAR5 were shown to be involved in cuticular wax primary alcohol biosynthesis in wheat anther and leaf cuticle (Y. Wang et al. 2015b, Y. Wang et al. 2015c). In addition, heterologous expression of the other three wheat TaFARs, TaFAR2, TaFAR3 and TaFAR4, in yeast (Saccharomyces cerevisiae) led to the accumulation of C18:0, C28:0 and C24:0 primary alcohols, respectively (M. Wang et al. 2016). Brachypodium distachyon (Brachypodium) has emerged as an excellent model system to facilitate biological investigations in grasses because of its small genome size, large collection of germplasm resources, short life cycle and simple growing conditions (Draper et al. 2001). The recent release of the genome sequence of the Bd21 accession further accelerates research in this model system (International Brachypodium Initiative 2010). Compared with C28 primary alcohol as the prevalent wax compound in wheat leaf blades (Y. Wang et al. 2015a, Y. Wang et al. 2015b, Y. Wang et al. 2015c), cuticular wax of six Brachypodium organs, i.e. cotyledons, spikes, leaf blades, leaf sheaths, stems and internal stems, was dominated by C26 primary alcohol (Luna 2014). Consequently, the identification of new C26 primary alcohol biosynthetic genes will further enhance our understanding of the substrate range of plant FARs, and this may provide a useful clue for enhancing the C26 primary alcohol production of wheat leaf blades by expressing C26 primary alcohol biosynthetic genes of Brachypodium in future wheat breeding programs. It will be very interesting to determine whether transgenic wheat lines containing high amounts of C26 and C28 primary alcohols could enhance drought tolerance compared with common wheat. In this study, we aimed to monitor the leaf wax profile and surface wax crystals of Brachypodium during leaf development, and to test whether three putative FAR genes, named BdFAR1, BdFAR2 and BdFAR3, are involved in the biosynthesis of cuticular wax primary alcohols for the Brachypodium cuticle. Results Kinetic changes in cuticular wax profiles of developing Brachypodium leaves Total wax was extracted from Brachypodium Bd21 leaves at five growth stages (20, 40, 60, 80 and 100 d).The total wax load increased from 3.77 μg cm−2 at 20 d to 4.85 μg cm−2 at 40 d, and then decreased steadily to 3.13, 2.37 and 1.05 μg cm−2 at 60, 80 and 100 d, respectively (Fig. 1A; Supplementary Table S1). Throughout leaf development, the wax mixture was dominated by primary alcohols (82.40–89.00%), first increasing to 4.32 μg cm−2 at day 40, and then continuously decreasing to 0.87 μg cm−2 at day 100 (Fig. 1A), suggesting that wax biosynthesis of leaves reached a peak at 40 d, especially primary alcohol biosynthesis. They were accompanied by alkanes (7.27–9.99%) and aldehydes (1.78–4.42%), which also increased from 20 to 40 d and then decreased between 60 and 100 d (Fig. 1A). In contrast, fatty acids were present in small amounts (0.94–3.19%) that decreased throughout the entire sampling period (Fig. 1A; Supplementary Table S1). Fig. 1 View largeDownload slide Cuticular wax accumulation and chain length distribution of the individual wax constituents on Bd21 leaves. (A) Developmental changes in the individual wax components and total load. (B) Developmental changes in chain length of the individual wax constituents. Numbers along the x-axis in (B) refer to the total carbon numbers of the compounds. Five representative developmental stages (20, 40, 60, 80 and 100 d) were investigated, and the amounts of cuticular wax are expressed as μg cm−2 of the leaf blade surface area. Each value is the average from three separate samples, and error bars indicate the SD. Fig. 1 View largeDownload slide Cuticular wax accumulation and chain length distribution of the individual wax constituents on Bd21 leaves. (A) Developmental changes in the individual wax components and total load. (B) Developmental changes in chain length of the individual wax constituents. Numbers along the x-axis in (B) refer to the total carbon numbers of the compounds. Five representative developmental stages (20, 40, 60, 80 and 100 d) were investigated, and the amounts of cuticular wax are expressed as μg cm−2 of the leaf blade surface area. Each value is the average from three separate samples, and error bars indicate the SD. The primary alcohols had predominantly even-numbered carbon chain lengths between C22 and C32, peaking at C26 throughout the growth period. Alkanes were found with mainly odd-numbered carbon chain lengths ranging from C25 to C33, with C29 alkane dominating between 20 and 40 d, and C29 and C31 alkanes were the major homologs from 60 to 100 d (Fig. 1B). The major aldehydes detected had C22 and C24 chains, while C26 and C28 aldehydes were present only in trace amounts that could not be quantified. Finally, the major fatty acid had a chain length of C16, accompanied by substantial amounts of C18 fatty acid (Fig. 1B) and traces of C20 and C22 fatty acids. Neither the aldehyde nor the fatty acid chain length profiles changed during leaf ontogenesis. Kinetic changes in wax crystal morphology of developing Brachypodium leaves To visualize the dynamic development of the epicuticular wax crystals on Brachypodium leaf blades, their adaxial and abaxial sides were investigated by SEM at the five growth stages as described above. At 20 d, both leaf sides were covered with very similar platelet-shaped wax crystals that were standing upright, with widely varying angles between them (Supplementary Fig. S1). The size of the wax platelets was typically 0.2–0.4 μm long and 0.1–0.2 μm high (Supplementary Fig. S1). At 40 d, platelet-shaped structures on both the adaxial and abaxial sides had increased lengths of 0.4–0.8 μm and heights of 0.2–0.4 μm, and were arranged in denser networks, suggesting increased numbers of crystals per unit area relative to 20 d. At 60 d, the crystal shapes and arrangements were similar to those at 40 d (Supplementary Fig. S1). During further leaf development, the networks of crystals on both leaf surfaces were gradually thinned, reflecting a decrease in the number of wax crystals compared with leaves at 60 d. Overall, the initial increase and final decrease of wax crystal numbers paralleled similar changes in the total wax load. Isolation and structure analysis of the Brachypodium BdFAR1, BdFAR2, and BdFAR3 genes Based on the finding that Brachypodium cuticular wax was dominated by primary alcohols throughout leaf development (Supplementary Table S1), we hypothesized that homologs of the Arabidopsis CER4 and wheat TaFARs may be important for Brachypodium cuticle formation. To identify FAR-like genes, the Brachypodium genome database was queried with the deduced amino acid sequence of wheat TaFAR1 (GenBank accession No. KF926683) using BLAST search programs. This revealed a group of six Brachypodium FAR-like sequences, of which the three most highly related to TaFAR1 were designated as BdFAR1, BdFAR2 and BdFAR3 (GenBank accession Nos. MF285084, MF285085 and MF285086, respectively). To isolate the BdFAR1, BdFAR2 and BdFAR3 coding regions, reverse transcription–PCR (RT–PCR) was performed using cDNA from Bd21 leaves. BdFAR1, BdFAR2 and BdFAR3 were amplified using the primer pairs BdFAR1-CDS, BdFAR2-CDS and BdFAR3-CDS, respectively, and all primer sequences are listed in Supplementary Table S2. The full-length cDNAs of BdFAR1, BdFAR2 and BdFAR3 are 1,904, 2,012 and 1,856 bp in length, comprising open reading frames (ORFs) of 1,584, 1,533 and 1,494 bp, respectively (Supplementary Fig. S2A–C). The genomic DNA (gDNA) sequence of BdFAR1 spans 3,817 bp with five exons and four introns encoding a 527 amino acid protein. The entire BdFAR2 exon–intron region is 3,931 bp in length and contains eight exons and seven introns encoding a 510 amino acid polypeptide. The gDNA sequence of BdFAR3 spans 4,482 bp, with 10 exons and nine introns encoding a 497 amino acid protein (Supplementary Fig. S2D). Analysis of the BdFAR protein sequences The predicted molecular masses of the BdFAR1, BdFAR2 and BdFAR3 proteins were 58.3, 57.3 and 55.9 kDa, respectively. This predicted result was confirmed by SDS–PAGE analysis of histidine-tagged BdFAR1, BdFAR2 and BdFAR3 proteins (Fig. 2A–C). A predictive protein analysis through the SMART website (http://smart.embl-heidelberg.de/) indicated that the three BdFARs all contained an NAD_binding_4 domain (NADB) at the N-terminus and a male-sterile domain at the C-terminus (Fig. 2D) (Aarts et al. 1997). Previous studies have shown that all plant FARs contained two conserved motifs, an NAD(P)H-binding site motif GXXGXX(G/A) and a classic YXXXK active site motif (Aarts et al. 1997), both of which were also observed in the BdFARs (Fig. 2D). Phylogenic analysis of the 18 plant FARs showed that they can be grouped into three distinct clades (Fig. 2E). The three BdFARs and five wheat TaFARs form the first clade, and seven dicot proteins were grouped into the second clade, including Artemisia annua GFAR1, jojoba ScFAR and Arabidopsis AtFAR1, AtFAR3, AtFAR4, AtFAR5 and AtFAR8. Additionally, the third clade contained rice DPW, Arabidopsis AtFAR2 and AtFAR6. Notably, the FARs in the first and second clades were predicted to be localized in the endoplasmic reticulum (ER), whereas FARs in the third clade were thought to be localized in plastids (Fig. 2E). Overall, the three BdFARs were closely related to the wheat TaFARs involved in primary alcohol biosynthesis for the wheat cuticle, suggesting that the BdFARs play an analogous role in wax alcohol formation in Brachypodium, and thus the biochemical functions of all three BdFARs were further analyzed in the present study. Fig. 2 View large Download slide Sequence analysis of BdFAR1, BdFAR2 and BdFAR3. (A–C) SDS–PAGE of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) proteins. Arrows show the HIS-BdFAR1, HIS-BdFAR2 and HIS-BdFAR3 proteins. (D) Sequence alignment of BdFAR1, BdFAR2 and BdFAR3 proteins. Physicochemically similar residues are shaded in gray. The predicted NAD_binding_4 and active sites are indicated in red boxes. The NAD_binding_4 domain and the male-sterile domain are indicated by black lines and stars under the sequences, respectively. (E) Phylogenetic analysis of plant FARs. This phylogenetic tree was constructed by MEGA 5.0 with the Neighbor–Joining method. The number for each interior branch refers to the percentage of the bootstrap value (1,000 replicates). The GenBank accession numbers of plant FAR genes used in the analysis are summarized in Supplementary Table S3. Fig. 2 View large Download slide Sequence analysis of BdFAR1, BdFAR2 and BdFAR3. (A–C) SDS–PAGE of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) proteins. Arrows show the HIS-BdFAR1, HIS-BdFAR2 and HIS-BdFAR3 proteins. (D) Sequence alignment of BdFAR1, BdFAR2 and BdFAR3 proteins. Physicochemically similar residues are shaded in gray. The predicted NAD_binding_4 and active sites are indicated in red boxes. The NAD_binding_4 domain and the male-sterile domain are indicated by black lines and stars under the sequences, respectively. (E) Phylogenetic analysis of plant FARs. This phylogenetic tree was constructed by MEGA 5.0 with the Neighbor–Joining method. The number for each interior branch refers to the percentage of the bootstrap value (1,000 replicates). The GenBank accession numbers of plant FAR genes used in the analysis are summarized in Supplementary Table S3. Heterologous expression of BdFARs in yeast To determine their biochemical activities and substrate specificities, the BdFAR1–BdFAR3 proteins were first expressed in yeast. To this end, the three coding regions were subcloned into the pYES2 vector for expression under the control of the yeast GAL1 promoter, and the recombinant plasmids were transformed into the yeast mutant strain INVSc1. Gas chromatography–mass spectroscopy (GC-MS) analysis showed that control yeast cells transformed with the empty vector accumulated C16:0, C16:1, C18:0 and C18:1 fatty acids, but no primary alcohols (Fig. 3A). In contrast, the expression of BdFAR1 resulted in the production of C22 alcohol (C22:0-OH), accompanied by small amounts of C24:0-OH (Fig. 3B). Similarly, the expression of BdFAR2 led to the formation of C24:0-OH and C26:0-OH (Fig. 3C), and the expression of BdFAR3 afforded C26:0-OH and minor amounts of C22:0-OH (Fig. 3D). The yeast expression experiments were repeated three times, consistently showing similar results. In summary, these results showed that the three BdFAR proteins have FAR activities to reduce fatty acyl-CoAs to primary alcohols without releasing aldehyde intermediates. The preferred substrate for BdFAR1 is C22:0 fatty acyl-CoA, while that of BdFAR2 and BdFAR3 is C26:0 fatty acyl-CoA, suggesting that both BdFAR2 and BdFAR3 may be involved in the formation of the major Brachypodium leaf wax component, C26 primary alcohol. Fig. 3 View largeDownload slide Heterologous expression of BdFAR1, BdFAR2 and BdFAR3 in yeast. Yeasts were transformed with empty vector control pYES2 (A) or with the pYES2 vector harboring BdFAR1 (B), BdFAR2 (C) or BdFAR3 (D). Major peaks were identified by GC-MS. In the empty vector control, fatty acids (16:1, 16:0, 18:1 and 18:0) but no primary alcohols were detected. In contrast, the yeast strains expressing BdFARs produced novel compounds identified as C22 primary alcohol (1-docosanol; C22:0-OH), C24 primary alcohol (1-tetracosanol; C24:0-OH) and C26 primary alcohol (1-hexacosanol; C26:0-OH). Fig. 3 View largeDownload slide Heterologous expression of BdFAR1, BdFAR2 and BdFAR3 in yeast. Yeasts were transformed with empty vector control pYES2 (A) or with the pYES2 vector harboring BdFAR1 (B), BdFAR2 (C) or BdFAR3 (D). Major peaks were identified by GC-MS. In the empty vector control, fatty acids (16:1, 16:0, 18:1 and 18:0) but no primary alcohols were detected. In contrast, the yeast strains expressing BdFARs produced novel compounds identified as C22 primary alcohol (1-docosanol; C22:0-OH), C24 primary alcohol (1-tetracosanol; C24:0-OH) and C26 primary alcohol (1-hexacosanol; C26:0-OH). Heterologous expression of BdFARs in transgenic tomato leaves and fruits To assess further the biochemical role of BdFAR in planta, we expressed the BdFAR coding regions in tomato (Solanum lycopersicum) cv MicroTom under the control of the Cauliflower mosaic virus (CaMV) 35S promoter via Agrobacterium infiltration (Supplementary Fig. S3A). The transgenic lines carrying the empty vector were used as control. Transgenic plants were screened using hygromycin selection and confirmed by PCR (Supplementary Fig. S3B). All of these transgenic lines displayed similar macroscopic growth phenotypes (Supplementary Fig. S3C). Compared with control plants, all the transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3 had significantly increased amounts of total primary alcohols, whereas the other wax components including n-alkanes, branched alkanes and triterpenoids were not significantly affected (Fig. 4A). In leaves of the transgenic BdFAR1-1 line, the C22:0-OH, C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH contents increased approximately 3.6-, 4.9-, 2.8-, 1.5- and 1.1-fold, respectively, while C32:0-OH was not affected. In the transgenic BdFAR2-2 line, the amounts of C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH increased approximately 6.9-, 4.0-, 2.0- and 1.5-fold, respectively, whereas the amounts of C22:0-OH and C32:0-OH remained nearly the same. Similarly, the transgenic BdFAR3-1 line showed 8.0-, 3.7-, 3.0- and 1.4-fold increases in C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH, respectively, while C22:0-OH and C32:0-OH were not significantly altered (Fig. 4C). Fig. 4 View largeDownload slide Cuticular wax accumulation in transgenic tomato cv MicroTom. (A, B) Cuticular wax amounts and compound class distribution in the wax of leaves (A) and ripe fruits (B) of T1 transgenic tomato lines. (C, D) Chain length distribution within the primary alcohols in the wax of leaves (C) and ripe fruits (D) of T1 transgenic lines. Control is the empty pCXSN vector plants. Other lines contain the coding regions of the BdFAR genes under the control of the 35S promoter. The data represent the means ± SD of three replicates. Significance is assessed by t-test (*P < 0.05, **P < 0.01). Fig. 4 View largeDownload slide Cuticular wax accumulation in transgenic tomato cv MicroTom. (A, B) Cuticular wax amounts and compound class distribution in the wax of leaves (A) and ripe fruits (B) of T1 transgenic tomato lines. (C, D) Chain length distribution within the primary alcohols in the wax of leaves (C) and ripe fruits (D) of T1 transgenic lines. Control is the empty pCXSN vector plants. Other lines contain the coding regions of the BdFAR genes under the control of the 35S promoter. The data represent the means ± SD of three replicates. Significance is assessed by t-test (*P < 0.05, **P < 0.01). Tomato fruits have wax distinct from that on leaves, with C26:0-OH, C28:0-OH, C30:0-OH and C32:0-OH as major alcohol components, thus enabling further comparative in planta investigations into the effects of BdFAR1–BdFAR3. We found that tomato fruits expressing BdFAR1, BdFAR2 or BdFAR3 had significantly increased amounts of primary alcohols (Fig. 4B), similar to corresponding transgenic leaves. In particular, all the transgenics expressing BdFARs showed increased amounts of C28:0-OH–C32:0-OH, while C34:0-OH remained unchanged relative to the empty vector control. Most interestingly, the lines expressing either BdFAR2 or BdFAR3 had markedly increased amounts of C26:0-OH (Fig. 4D). In addition, we did not observe significant differences in the crystals on leaves and fruits between all transgenic lines and control plants (Supplementary Fig. S4). Taken together, the tomato expression results confirmed that the tested Brachypodium enzymes are active FARs, which accept fairly broad ranges of C24–C32 acyl-CoA substrates. BdFAR2 and BdFAR3 were characterized by relatively strong activity on C26 fatty acyl-CoA, while BdFAR1was distinguished by its ability also to utilize the C22 fatty acyl-CoA substrate. Heterologous expression of BdFARs in Arabidopsis cer4-3 mutant leaves As our yeast and tomato expression results partially diverged, we sought to test further the biochemical characteristics of the BdFAR proteins by heterologous expression in another plant system. To this end, we expressed the BdFAR genes in the Arabidopsis cer4-3 mutant deficient in the formation of C22–C28 primary alcohols (Rowland et al. 2006) and analyzed the wax composition of leaves expressing the BdFAR genes in comparison with empty vector controls. The total amounts of primary alcohols in the BdFAR1 and BdFAR3 transgenic lines were very close to those of the control lines, while the primary alcohol coverages on the BdFAR2 transgenic lines were dramatically increased compared with empty vector control plants (Fig. 5A). All other wax components, including fatty acids, aldehydes, branched alcohols, alkanes, ketone, secondary alcohols and sterols, did not change significantly between the BdFAR transgenic lines and the empty vector control (Fig. 5A). Fig. 5 View largeDownload slide Cuticular wax analysis on leaves of the transgenic Arabidopsis cer4-3 mutant. (A) Distribution of individual compound classes in wax of leaves of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. (B) Chain length distribution within the primary alcohols in the leaf wax of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. Six-week-old Arabidopsis leaves were harvested for cuticular wax analysis. The amounts of wax components are expressed as μg cm−2 leaf surface area. The data represent the means ± SD of three replicates. Asterisks indicate significant differences between the empty vector control and BdFAR1, BdFAR2 or BdFAR3 transgenic lines according to t-test (*P < 0.05; **P < 0.01). Fig. 5 View largeDownload slide Cuticular wax analysis on leaves of the transgenic Arabidopsis cer4-3 mutant. (A) Distribution of individual compound classes in wax of leaves of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. (B) Chain length distribution within the primary alcohols in the leaf wax of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. Six-week-old Arabidopsis leaves were harvested for cuticular wax analysis. The amounts of wax components are expressed as μg cm−2 leaf surface area. The data represent the means ± SD of three replicates. Asterisks indicate significant differences between the empty vector control and BdFAR1, BdFAR2 or BdFAR3 transgenic lines according to t-test (*P < 0.05; **P < 0.01). Within the primary alcohol fraction in leaf wax of Arabidopsis expressing BdFAR1, the amounts of C22:0-OH, C24:0-OH and C26:0-OH significantly increased, while the amounts of C28:0–C34:0 alcohol did not differ compared with empty vector control plants (Fig. 5B). In the transgenic line expressing BdFAR2, a sharp increase in the content of C26:0-OH was observed along with slightly increased levels of C24:0-OH and C28:0-OH compared with the control (Fig. 5B). Moreover, the expression of BdFAR3 led to increased levels of C22:0-OH, C24:0-OH and C26:0-OH, along with a slight decrease of C32:0-OH relative to control plants, while primary alcohols with other carbon lengths did not show significant changes (Fig. 5B). Together, these results further confirm that BdFARs are active enzymes involved in wax formation in Brachypodium, further suggesting that BdFAR1 and BdFAR2 have preferences for C22 and C26 substrates, respectively. Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves To examine whether BdFAR is involved in the formation of wax alcohols in Brachypodium, we constitutively overexpressed both genes under the control of the CaMV 35S promoter. To this end, the binary vector constructs pCXSN-BdFARs were transformed into Brachypodium (accession Bd21). Unfortunately, our attempt to obtain pCXSN-BdFAR1 transgenic plants was unsuccessful. A total of six independent transgenic BdFAR2 and BdFAR3 lines were obtained. In BdFAR2 and BdFAR3 overexpression lines, the transcript level of BdFAR2 and BdFAR3 significantly increased compared with that in the wild-type (WT) plants (Fig. 6A). The total amounts of alkanes, fatty acids and aldehydes were not affected in the different overexpression lines compared with WT plants, whereas the amounts of primary alcohols were significantly higher in all the overexpression lines (Fig. 6B). In particular, all transgenic lines had levels of C26 alcohol significantly increased relative to the WT, and the amounts of C28 alcohol were also increased in most lines (Fig. 6C). In contrast, the other wax alcohols (C22:0-OH, C24:0-OH, C30:0-OH and C32:0-OH) did not show significant changes between overexpression lines and WT plants (Fig. 6C). Additionally, there were no significant differences in wax crystals on leaves between all transgenic lines and WT plants (Supplementary Fig. S5). Most interestingly, these findings confirmed some but not all of the details in our results from the heterologous expression of the same genes in yeast and other plants. Overall, these results clearly indicated that BdFAR2 and BdFAR3 play a key role in wax alcohol production in Brachypodium, especially the C26 alcohol, which is the major wax component of Brachypodium leaf wax. Fig. 6 View largeDownload slide Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves. (A) qRT-PCR analysis of BdFAR2 and BdFAR3 in WT plants, and BdFAR2- and BdFAR3-overexpressing lines. (B) Distribution of individual compound classes in wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. (C) Chain length distribution within the primary alcohols in the leaf wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. Error bars represent the SD (n =3). The data were statistically analyzed using t-test (*P < 0.05, **P < 0.01). Fig. 6 View largeDownload slide Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves. (A) qRT-PCR analysis of BdFAR2 and BdFAR3 in WT plants, and BdFAR2- and BdFAR3-overexpressing lines. (B) Distribution of individual compound classes in wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. (C) Chain length distribution within the primary alcohols in the leaf wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. Error bars represent the SD (n =3). The data were statistically analyzed using t-test (*P < 0.05, **P < 0.01). Temporal and spatial patterns of BdFAR genes in Brachypodium To better understand the functions of BdFAR genes, we examined their spatial and temporal expression patterns in different vegetative and reproductive organs of Bd21. Quantitative real-time PCR (qRT-PCR) showed that the three BdFAR genes were expressed in aerial vegetative and reproductive organs, but not in roots. BdFAR1 was found to be highly expressed in early developing leaves, leaf sheaths, nodes and internodes, modestly expressed in late developing leaves, spikelets and glumes, but not expressed in roots and leaves at 100 d (Fig. 7A). BdFAR2 was mainly expressed in leaf sheaths, nodes, internodes and early developing leaves, and at very low levels in late developing leaves, spikelets and glumes, but not in roots (Fig. 7B). Similarly, the BdFAR3 transcript was detected at high levels in leaves at 40 d, leaf sheaths and internodes, at lower levels in nodes, leaves at 20, 60 and 80 d, spikelets and glumes, but not in roots and leaves at 100 d (Fig. 7C). Notably, the expression of all three BdFAR genes increased greatly between 20 and 40 d, and then gradually decreased during further leaf development, thus paralleling the changes in amounts of primary alcohols on leaf surfaces during leaf development. Fig. 7 View largeDownload slide Temporal and spatial expression patterns of the Brachypodium BdFAR1, BdFAR2 and BdFAR3 genes. (A) Differential expression analysis of BdFAR genes in various organs of Brachypodium by qRT-PCR. LB20, LB40, LB60, LB80 and LB100 represent leaf blades at 20, 40, 60, 80 and 100 d of plant development, respectively. The Brachypodium UBI4 gene was used to normalize gene expression, and error bars represent ± SD of three biological replicates. (B–I) GUS staining analysis of the BdFAR1 (B–E) and the BdFAR2 (F–I) promoter activities in transgenic Brachypodium plants. (B, F) Leaf blades of 50-day-old plants, (C, G) leaf sheaths, (D, H) internodes, (E, I) nodes. Scale bars = 1 mm. Fig. 7 View largeDownload slide Temporal and spatial expression patterns of the Brachypodium BdFAR1, BdFAR2 and BdFAR3 genes. (A) Differential expression analysis of BdFAR genes in various organs of Brachypodium by qRT-PCR. LB20, LB40, LB60, LB80 and LB100 represent leaf blades at 20, 40, 60, 80 and 100 d of plant development, respectively. The Brachypodium UBI4 gene was used to normalize gene expression, and error bars represent ± SD of three biological replicates. (B–I) GUS staining analysis of the BdFAR1 (B–E) and the BdFAR2 (F–I) promoter activities in transgenic Brachypodium plants. (B, F) Leaf blades of 50-day-old plants, (C, G) leaf sheaths, (D, H) internodes, (E, I) nodes. Scale bars = 1 mm. To investigate further the gene expression patterns of BdFAR1 and BdFAR2, genomic promoter sequences 2,264 and 2,285 bp upstream of the start codons, respectively, were fused to the β-glucuronidase (GUS) reporter gene. For both promoters, strong GUS activity was detected in leaf blades (Fig. 7B, F), leaf sheaths (Fig. 7C, G) and nodes (Fig. 7E, I), while only modest GUS activity was observed in internodes (Fig. 7D, H). Unfortunately, our attempt to obtain pBdFAR3:GUS transgenic plants was unsuccessful. Overall, the GUS results are thus consistent with the qRT-PCR data. The expression patterns support the idea that all three BdFARs play a role in the production of primary alcohols for the cuticular wax in Brachypodium. Subcellular localization of BdFAR proteins In silico analyses predicted BdFAR to be localized in the ER (Fig. 2E). To localize the three BdFARs, the corresponding full-length coding sequences without stop codons were fused in-frame to the N-terminus of the green fluorescent protein (GFP) gene for expression under the control of the CaMV 35S promoter, and the resulting constructs were introduced into Arabidopsis leaf protoplasts using polyethylene glycol- (PEG) mediated transformation, and a sequence encoding the ER-specific protein mCherry-HDEL fused with red fluorescent protein (RFP) was co-transformed into the protoplasts (Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016). Both the green BdFAR–GFP and the red HDEL–RFP signals were visualized by confocal microscopy, showing that the GFP fusions of all three BdFARs co-localized completely with the mCherry-HDEL signals (Fig. 8). Consequently, the BdFAR proteins were localized to the ER; thus, the subcellular localization experiments completely confirm the results of the previous in silico analysis (Fig. 2E). Fig. 8 View largeDownload slide Subcellular localization of BdFAR1, BdFAR2 and BdFAR3 in Arabidopsis leaf protoplasts. Each row of five images shows, from left to right, the GFP signal of the BdFAR–GFP fusion construct, the RFP signal of the ER marker mCherry-HDEL, blue fluorescence from the Chl autofluorescence signal, bright-field image and merge of the previous four images. Bars = 5 μm. Fig. 8 View largeDownload slide Subcellular localization of BdFAR1, BdFAR2 and BdFAR3 in Arabidopsis leaf protoplasts. Each row of five images shows, from left to right, the GFP signal of the BdFAR–GFP fusion construct, the RFP signal of the ER marker mCherry-HDEL, blue fluorescence from the Chl autofluorescence signal, bright-field image and merge of the previous four images. Bars = 5 μm. Transcriptional regulation of BdFAR genes under abiotic stress Recent evidence indicated that wax accumulation on the aerial surfaces of diverse plant species could be modulated by water deficiency, cold and ABA treatments (W. Wang et al. 2015a, Y. Wang et al. 2015c, M. Wang et al. 2016). To investigate whether BdFAR expression is regulated by abiotic stresses, we monitored the BdFAR transcript levels in 60-day-old Bd21 seedlings under stress for up to 24 h. When seedlings were subjected to drought, transcript levels of BdFAR1 and BdFAR2 increased approximately 23- and 8-fold, respectively, from the start of the treatment until 24 h (Fig. 9A, B). In contrast, BdFAR3 expression peaked at 2 h, showing a 9-fold increase compared with 0 h (Fig. 9C). Consistent with the drought induction, PEG treatment resulted in approximately 11-, 12- and 9-fold increases in BdFAR1, BdFAR2 and BdFAR3 transcript levels, respectively (Fig. 9A–C). Likewise, cold treatment led to 6-, 4- and 3-fold increases in BdFAR1, BdFAR2 and BdFAR3, respectively (Fig. 9A–C). Based on these observations, we concluded that transcription of the BdFAR genes is positively regulated by abiotic stresses including water deficiency and cold treatments. Next, we examined whether the three BdFAR genes are influenced by the phytohormone ABA. Treatment with 100 μM ABA led to 5-, 23- and 6-fold increases in BdFAR1, BdFAR2 and BdFAR3 transcript abundances, respectively (Fig. 9A–C). From this finding, we concluded that the phytohormone ABA plays an important role in the transcriptional control of BdFAR1–BdFAR3 activation. Fig. 9 View largeDownload slide Expression analysis of BdFAR1, BdFAR2 and BdFAR3 under abiotic stress and ABA treatments. Time course of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) expression after abiotic stress treatment and ABA application. Sixty-day-old soil-grown plants were exposed to various stresses, and relative transcript levels were quantified by qRT-PCR using BdUBI4 as a reference for normalization of the sample amounts. For drought treatment, plants were removed from the soil and allowed to dry under 60% humidity; PEG treatment, 20% (w/v) PEG 6000; Cold treatment, 4°C; ABA treatment, 100 μM ABA. (D) Total wax amounts between well-watered plants (Bd21) and plants after 10 d of water deprivation (Bd21/DR). (E) Cuticular wax composition on leaves of control Bd21 and Bd21/DR plants. Each value represents the mean of three independent measurements. Error bars indicate the SD. Data were statistically analyzed using t-test (*P < 0.05; **P < 0.01). Fig. 9 View largeDownload slide Expression analysis of BdFAR1, BdFAR2 and BdFAR3 under abiotic stress and ABA treatments. Time course of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) expression after abiotic stress treatment and ABA application. Sixty-day-old soil-grown plants were exposed to various stresses, and relative transcript levels were quantified by qRT-PCR using BdUBI4 as a reference for normalization of the sample amounts. For drought treatment, plants were removed from the soil and allowed to dry under 60% humidity; PEG treatment, 20% (w/v) PEG 6000; Cold treatment, 4°C; ABA treatment, 100 μM ABA. (D) Total wax amounts between well-watered plants (Bd21) and plants after 10 d of water deprivation (Bd21/DR). (E) Cuticular wax composition on leaves of control Bd21 and Bd21/DR plants. Each value represents the mean of three independent measurements. Error bars indicate the SD. Data were statistically analyzed using t-test (*P < 0.05; **P < 0.01). Studies have shown that the stress-induced expression of wax biosynthesis genes may lead to altered amounts of wax and composition in several plant species (Shepherd and Wynne Griffiths 2006, Panikashvili et al. 2007, Joubès et al. 2008, Kosma et al. 2009). Therefore, we next tested whether the drought effects on BdFAR1–BdFAR3 expression were paralleled by changes in the composition of Brachypodium leaf wax. To this end, we extracted wax from well-irrigated plants (Bd21) and drought-treated plants (Bd21/DR), followed by analysis using GC-MS and gas chromatography with flame ionization detection (GC-FID). After approximately 10 d of drought treatment, leaves of Bd21/DR plants began to wilt (Supplementary Fig. S6). Compared with Bd21, Bd21/DR exhibited a significant increase in the total amount of leaf wax (Fig. 9D), with a 90% increase in fatty acids, an 80% increase in aldehydes, a 60% increase in alkanes and a 60% increase in primary alcohols (Fig. 9E). However, there were no significant shifts in the proportion of these compound classes, or in the chain length distributions within them between Bd21 and Bd21/DR. Altogether, our observations indicate that drought stress induces cuticular wax accumulation in Brachypodium leaves, and that the BdFAR1–BdFAR3 genes play a role in the drought-induced accumulation of cuticular wax, particularly wax alcohols. Discussion Here, we systematically investigated the developmental changes in cuticular wax load, composition and morphology during leaf development in Brachypodium. Our data indicated that primary alcohols were the major components of cuticular wax on Brachypodium leaf blades throughout the entire period of plant growth. Then, we functionally characterized three Brachypodium FAR genes, and showed their involvement in the biosynthesis of cuticular wax primary alcohols. The amounts and crystal densities of cuticular wax vary at different 0developmental stages in Brachypodium In the present study, we observed dynamic changes in wax coverage on Brachypodium leaves, with initially increasing and then continuously decreasing amounts during leaf development (Fig. 1A; Supplementary Table S1). To our knowledge, the present investigation for the first time reports developmental changes of cuticular wax in Brachypodium. Developmental changes in the amount and composition of wax have also been observed in other diverse crops (Peschel et al. 2007, Y. Wang et al. 2015a). The cuticular wax of Brachypodium consisted of the four compound classes, namely primary alcohols, alkanes, aldehydes and fatty acids, and primary alcohols dominated the wax mixture throughout leaf development, as well as primary alcohols being the dominant component in six Brachypodium organs (Luna 2014). We hypothesize that the acyl reduction pathway may play a key role in wax biosynthesis of Brachypodium cuticle. Notably, hexacosanol was the most abundant primary alcohol on Brachypodium leaf surfaces, whereas octacosanol was the predominant primary alcohols on wheat leaves (Adamski et al. 2013, Zhang et al. 2013). In addition, diketone, a key wax component in wheat, barley (Hordeurn vulgare L.) and Arabidopsis (Greer et al. 2007, Y. Wang et al. 2015b, Schneider et al. 2016), has not been detected, suggesting that the glaucous character may be not be present in Brachypodium cuticle. Throughout Brachypodium leaf development, both the adaxial and abaxial leaf surfaces displayed similar platelet-shaped wax crystals (Supplementary Fig. S1). Interestingly, the sizes of the platelets increased, and their arrangements first increased in density then gradually decreased, thus matching the changes in the total amounts of wax. Therefore, our results further support the notion that the formation of platelet crystals is positively correlated with primary alcohols on Brachypodium leaf surfaces. This finding is consistent with previous reports on wax platelets formed by alcohols in other plants (Koch et al. 2006, Zhang et al. 2013, Y. Wang et al. 2015c). Taken together, our findings suggest that differences in surface morphology are driven by dynamic changes in wax alcohol accumulation and underlying quantitative changes in the biosynthesis of the wax alcohols. BdFAR1, BdFAR2 and BdFAR3 are involved in the formation of primary alcohols for the Brachypodium cuticle Here, we isolated three BdFAR genes from Brachypodium. To our knowledge, the BdFAR genes are the first in Brachypodium to be characterized in detail with regard to primary alcohol biosynthesis. Transgenic expression of BdFARs in yeast, tomato, the Arabidopsis cer4-3 mutant and Brachypodium afforded production of primary alcohols, indicating that three BdFAR proteins have FAR activities catalyzing the reduction of fatty acyl-CoAs to primary alcohols. In particular, overexpression of BdFAR2 and BdFAR3 strongly enhanced the amounts of both C26:0 and C28:0 primary alcohol in Brachypodium leaves, suggesting that BdFARs are a promising tool for future studies aiming at enhancing the amounts of plant wax. It is likely that the expression of BdFAR2 and BdFAR3 in wheat may result in high contents of both C26 and C28 primary alcohols and increase drought tolerance. Heterologous expression of yeast indicated that BdFAR1 prefers C22 acyl-CoA as the substrate, whereas BdFAR2 and BdFAR3 preferentially accept C26 acyl-CoA. In fact, BdFAR1 exhibits the highest sequence identity to that of TaFAR1 (75%) and TaFAR5 (74%), which also use C22 acyl-CoA as the preferred substrate in yeast (Y. Wang et al. 2015b, Y. Wang et al. 2015c). Likewise, BdFAR3 shares 84% identity with TaFAR3, which prefers C26 acyl-CoA as the substrate in yeast (M. Wang et al. 2016). However, BdFAR2, TaFAR2 and TaFAR4 are highly similar, but they possess distinct specificities for C26, C24 and C18 acyl-CoAs, respectively (M. Wang et al. 2016). Consequently, it was concluded that sequence similarities of FARs were not correlated with their substrate specificities. The substrate preferences of the BdFAR1, BdFAR2 and BdFAR3 proteins are similar to those of homologous FAR enzymes in other species. For example, Arabidopsis AtFAR1 and AtFAR4 showed preferences for C22 and C20 acyl-CoAs, respectively (Domergue et al. 2010), whereas AtCER4 catalyzes the reduction of C24–C30 acyl-CoAs (Rowland et al. 2006). Jojoba ScFAR uses C20 and C22 acyl-CoAs as substrates (Metz et al. 2000) (Fig. 10). Fig. 10 View largeDownload slide Substrate range of plant FARs involved in synthesis of wax primary alcohols. CER6, KCR1, PAS2, CER10, FAR1, FAR4 and CER4 are wax biosynthetic enzymes from Arabidopsis. ScFAR is the jojoba FAR enzyme involved in the formation of seed storage wax esters. TaFAR1, TaFAR2, TaFAR3, TaFAR4 and TaFAR5 are the FARs involved in wax biosynthesis in wheat. BdFAR1, BdFAR2 and BdFAR3 are the FARs involved in wax biosynthesis in Brachypodium. LACS, long chain acyl-CoA synthetase; FAE, fatty acid elongase. Fig. 10 View largeDownload slide Substrate range of plant FARs involved in synthesis of wax primary alcohols. CER6, KCR1, PAS2, CER10, FAR1, FAR4 and CER4 are wax biosynthetic enzymes from Arabidopsis. ScFAR is the jojoba FAR enzyme involved in the formation of seed storage wax esters. TaFAR1, TaFAR2, TaFAR3, TaFAR4 and TaFAR5 are the FARs involved in wax biosynthesis in wheat. BdFAR1, BdFAR2 and BdFAR3 are the FARs involved in wax biosynthesis in Brachypodium. LACS, long chain acyl-CoA synthetase; FAE, fatty acid elongase. BdFARs are localized to the ER and preferentially expressed in aerial tissues Confocal microscopy observation revealed that BdFAR1, BdFAR2 and BdFAR3 were all localized to the ER (Fig. 8), the subcellular compartment known to harbor all wax biosynthesis reactions (Rowland et al. 2006, Greer et al. 2007, Mao et al. 2012, Rajangam et al. 2013, W. Wang et al. 2015a, Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016, Hegebarth et al. 2017). In Arabidopsis, both WSD1 catalyzing the terminal step of the primary alcohol pathway and the midchain hydroxylase MAH1 catalyzing the final step of the alkane pathway were present in the ER (Greer et al. 2007, Li et al. 2008), leading us to hypothesize that the ER is the core subcellular compartment where almost all cuticular wax components are generated in planta. Further experiments showed that the three investigated BdFAR genes were expressed in all the aerial parts of Brachypodium plants (Fig. 7), consistent with the putative role of these genes in wax production. In particular, the changes in BdFAR expression levels in the course of leaf development were in good accordance with the dynamic changes in the amounts of wax alcohols, further suggesting that the three BdFAR genes are the key enzymes dedicated to wax primary alcohol biosynthesis in Brachypodium. The preferential expression of the BdFARs in aerial organs of Brachypodium is similar to that of many wax biosynthesis genes in other plant species, including Arabidopsis and wheat (Rowland et al. 2006, Greer et al. 2007, Bourdenx et al. 2011, Haslam et al. 2012, Y. Wang et al. 2015c, Y. Wang et al. 2015c, M. Wang et al. 2016). Conversely, it is interesting to note that the three BdFAR genes were barely expressed in roots, suggesting that they are not involved in the formation of suberin or suberin-associated wax. This is in contrast to some of the FARs of Arabidopsis, where AtFAR1, AtFAR4 and AtFAR5 were implicated in suberin biosynthesis (Domergue et al. 2010), along with elongation enzymes encoded, for example, by the ketoacyl-CoA synthase genes KCS2, KCS20 and KCS9 (Franke et al. 2009, Lee et al. 2009, Kim et al. 2013). BdFAR gene expression is regulated by drought, cold stress and ABA treatment Surface wax mixtures play a critical role in protecting plants against biotic and abiotic stresses. One major function of cuticular wax is to prevent non-stomatal water loss, which is especially important for plant survival in water-limited environments (Kosma and Jenks 2007). Accordingly, it has been proposed that during severe water deficit, when stomata are closed, nanoscale diffusion pathways traversing the cuticle become the primary conduit of plant water loss (Kosma et al. 2009). It therefore seems plausible that plants may react to drought stress by modification of their cuticular wax composition, and water deficit may enhance wax synthesis by up-regulating the production of key wax biosynthetic enzymes (Hooker et al. 2002). Our results, showing that expression of the three BdFAR genes was induced under drought stress conditions, clearly support this hypothesis. In addition, BdFAR1 and BdFAR2 were dramatically up-regulated approximately 23- and 8-fold after 24 h of drought stress treatment, and BdFAR3 expression showed a 9-fold increase after 2 h of drought stress treatment, suggesting that induction of expression of BdFAR3 was faster than that of BdFAR1 and BdFAR2 under drought stress. Notably, Brachypodium plants also exhibited a significant increase in the amounts of total leaf wax under drought stress, thus further suggesting that the induction of BdFAR expression indeed leads to changes in the composition of cuticular wax similar to other diverse plant species (Kim et al. 2007a, Kim et al. 2007b, Kosma et al. 2009, Zhu and Xiong, 2013, Y. Wang et al. 2015b). How such changes in cuticle composition may alter its physiological properties and, ultimately, plant performance under drought stress is an important question for future study. Interestingly, ABA treatment also induced BdFAR expression. In Arabidopsis, ABA is known to induce the MYB96 transcription factor which directly binds to conserved sequence motifs in several wax biosynthesis gene promoters and modulates wax biosynthesis in response to drought (Seo et al. 2011). Therefore, it may be speculated that ABA regulation of wax biosynthesis in Brachypodium involves similar transcriptional control mechanisms to those in Arabidopsis, possibly also involving transcription factors such as MYB96. In this context, it should be noted that transcript levels of the BdFAR genes were up-regulated within a few hours under abiotic stresses and ABA treatments, demonstrating that wax production can change rapidly in response to stress. Consistent with our results, the abundances of Arabidopsis CER1 and wheat TaFAR transcripts also increased rapidly under abiotic stress (Bourdenx et al. 2011, Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016). In addition, BdFAR expression was induced by cold treatments, similar to the five wheat TaFAR genes (Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016), cucumber CsWAX2 as well as CsCER1 (W. Wang et al. 2015a, W. Wang et al. 2015b) and Medicago sativa WXP1 (Zhang et al. 2005) which were also up-regulated by cold treatment. These findings are in contrast to Arabidopsis, where cold stress has been found to down-regulate the expression of several components of the FAE complex that are crucial for wax biosynthesis (Joubès et al. 2008). It is likely that the cold stress response varied in severity and effect depending on the plant species. Materials and Methods Plant materials, growth conditions and stress treatments The Brachypodium genotype Bd21 was used for all experiments. Plants were grown in a glasshouse with standard irrigation and fertilization under long-day conditions (18 h of light). Day and night temperatures were 22 and 18°C, respectively. To analyze developmental changes in the wax load, composition and morphology, the first leaves were sampled at 20, 40, 60, 80 and 100 d after germination. Three leaves were used in one extraction and three replicates per stage were tested. Approximately 60 d after germination, Brachypodium plants were subjected to various stress treatments. For drought treatment, plants were air-dried on filter paper and harvested at different time points. For PEG and ABA treatments, plants were transferred to solutions containing 20% (w/v) PEG 6000 and 100 μM ABA, respectively. For cold stress treatment, plants were maintained at 4°C. Cuticular wax extraction and chemical analysis Brachypodium leaves were immersed in 50 ml of chloroform and shaken for 1 min at room temperature. A 20 μg aliquot of n-tetracosane was added as an internal standard. Solvent was evaporated from the resulting extracts under nitrogen, the samples were taken up in chloroform and transferred to a GC autosampler vial, dried again under nitrogen, and derivatized in 100 μl of pyridine and 100 μl of bis-(N, N-trimethylsilyl)-trifluoroacetamide (BSTFA) for 60 min at 70°C. Solvent was again removed under nitrogen and samples were dissolved in 500 μl of chloroform for analyses using GC-MS and GC-FID. The wax composition was determined using a capillary gas chromatograph equipped with an Rxi-5ms column (30 m length, i.d. 0.25 mm, df 0.25 μm; Restek) and a mass spectrometer (GCMS-QP2010, Shimadzu) with helium as the carrier gas. The oven program ran at 50°C for 2 min, increased temperature by 20°C min−1 to 200°C and held for 1 min, increased by 2°C min−1 to 320°C and held for 15 min. Injector and detector temperatures were set at 250°C, and 1 μl of each sample was injected. Identification of wax compounds was based on a comparison of their mass spectra with those of authentic standards and literature data. For the quantification of individual compounds, GC-FID (GC-2010Plus, Shimadzu) with an Rtx-1column (60 m length, i.d. 0.32 mm, df 0.25 μm; Restek) was used under the conditions described above, but with H2 carrier gas. Amounts of all detected compounds were assessed by comparison with peak areas relative to the internal standard, and wax coverage was calculated relative to the extracted surface area calculated by analysis of a digital photograph using Image J software. SEM Fresh leaves were collected from Bd21 plants at 20, 40, 60, 80 and 100 d after germination, air-dried for 10 d in a desiccator at room temperature, and then carefully dissected. Small pieces of samples were mounted on specimen stubs using double-sided copper tape, and sputtered with gold particles at 25 mA for 90 s in a Bal-Tec SCD005 sputter coater (Balzers). The coated surfaces were investigated by SEM (Hitachi S4800) at an accelerating voltage of 10 kV and a working distance of 12 mm. Cloning of BdFAR1, BdFAR2 and BdFAR3 Total RNA was extracted from Bd21 leaves using Trizol Reagent (Invitrogen). Purified RNA was treated with DNase I (Promega) to remove residual DNA, and then was used as the template for first-strand cDNA synthesis using PrimeScript™ reverse transcriptase (TAKARA). The BdFAR cDNAs were amplified with specific primers (Supplementary Table S2) under the following PCR conditions: 95°C for 5 min, 35 cycles of 95°C for 30 s, 58°C for 30 s and 72°C for 2 min, with a final extension at 72°C for 1 min. The amplified products were cloned into the pMD™ 18-T vector (TAKARA) using T4 DNA ligase (TAKARA) and sequenced for verification. The genomic sequences of BdFAR genes were obtained from Brachypodium gDNA. Heterologous expression in yeast The coding sequences of BdFAR genes were amplified from Bd21 cDNA using specific primers (labeled as BdFARx-YS listed in Supplementary Table S2), and the PCR products were cloned into the yeast expression vector pYES2 (Invitrogen) using the In-Fusion® HD Cloning Kit (Clontech). The resulting pYES-BdFARx constructs were transformed into Escherichia coli Top10 cells, and verified by colony PCR and sequencing. Subsequently, the pYES-BdFARx and an empty vector were transformed into the mutant yeast strain INVSc1 according to Gietz and Woods (2002). Transgenic yeast cells were grown on synthetic complete (SC) selection medium without uracil. Three individual cell lines were selected from each transgenic strain. First, yeast cells were inoculated into 20 ml of SC medium containing 2% glucose and grown for 2 d at 30°C, with shaking at 200 r.p.m. Yeast cells were then transferred to 20 ml of induction medium (SC medium containing 2% galactose) for 12 h at 30°C, with shaking at 200 r.p.m. The yeast cells were then transferred to 20 ml of resting medium (0.1 M potassium phosphate containing glucose and hemin) for 24 h at 30°C, with shaking at 200 r.p.m. Then, yeast cells were collected by centrifugation, refluxed for 5 min in 20% KOH/50% ethanol and extracted twice with hexane. Both hexane solutions were combined, and the solvent was removed under a gentle stream of nitrogen. The remaining procedure was consistent with the plant wax analysis described above. Tomato transformation For constitutive tomato expression under the control of the 35S promoter, the coding sequences for BdFAR genes were inserted into a binary vector pCXSN digested with XcmI (NEB) restriction enzyme (Chen et al. 2009). The resulting constructs and an empty vector were individually transformed into Agrobacterium strain GV3101. Cotyledon transformation of tomato cv MicroTom was performed as previously described (Dan et al. 2006). The presence of the transgene was confirmed in the T0 generation by PCR. Transgenic T0 and T1 generation plants were grown in the greenhouse (22°C with 8 h dark/16 h light). Wax analysis of mature leaves and mature red fruits from transgenic plants was performed as previously described (Wang et al. 2011). Arabidopsis transformation The constructs used for tomato transformation (see above) were introduced into the Arabidopsis cer4-3 mutant via Agrobacterium-mediated transformation using the floral dip method (Clough and Bent 1998). The transformed plants were screened on Murashige and Skoog (MS) medium with 25 mg l−1 hygromycin (w/v), and 6-week-old leaves of T1 plants were harvested for cuticular wax analysis. Brachypodium overexpression For BdFAR overexpression under the control of the 35S promoter, the BdFAR2 and BdFAR3 coding regions were amplified and inserted into the XcmI (NEB) site of the pCXSN vector. Then, the resulting constructs were introduced into Agrobacterium strain EHA105, and Brachypodium transformation was performed according to Vogel and Hill (2008). Kanamycin-resistant plants were screened by PCR for transgenes, and selected plantlets were transferred to soil, placed under a plastic wrap for approximately 1 week to acclimate the plants and then grown in the greenhouse. Sequence alignment and phylogenetic analysis The homologous BdFAR proteins were identified in the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov) via BLAST search. Multiple protein sequence alignments were performed with the ClustalW 1.83 program (Thompson et al. 1997), and the aligned sequences were imported into Bioedit for manual editing (http://www.mbio.ncsu.edu/BioEdit/bioedit.html; Hall 1999). A phylogenetic tree was constructed using the Neighbor–Joining (NJ) method by MEGA 5.0 software with the option of pairwise deletion, and 1,000 bootstrap replicates to test inferred phylogeny (Tamura et al. 2011). qRT-PCR For tissue expression analysis, leaf blades from plants at 20, 40, 60, 80 and 100 d, nodes, internodes, leaf sheaths, spikelets, glumes of 9-week-old plants, roots of 2-week-old plants and 60-day-old seedlings treated with various stresses were collected and ground in liquid nitrogen. Total RNA extractions were performed with Trizol Reagent (Invitrogen). After treatment with DNase (Promega), total RNA was used for the first-strand cDNA synthesis using PrimeScript™ reverse transcriptase (TAKARA). qRT-PCR was performed in a 25 μl volume using the SYBR® Premix Ex Taq™ Kit (TAKARA) on a CFX96 real-time PCR detection system (Bio-Rad). BdUBI4 (LOC100832168) was used for normalization of the amount of sample. Each experiment had three biological parallels, each with three technical replicates. Histochemical β-glucuronidase analysis For promoter analysis, the 2,264 and 2,285 bp genomic fragments located upstream of the ATG start codon of BdFAR1 and BdFAR2 were amplified and inserted in front of the GUS gene in pCAMBIA1305 using the In-Fusion® HD Cloning Kit (Clontech). The GUS fusion constructs pBdFAR1:GUS and pBdFAR2:GUS were then transformed into Bd21 plants using Agrobacterium (Vogel and Hill 2008). Histochemical GUS staining was performed as previously described (Jefferson et al. 1987). Transgenic plant samples were incubated in X-gluc buffer at 37°C overnight, and then the staining solution was removed and the samples were cleared of Chl with 70% ethanol. The stained tissues were observed and photographed using a SZX16 stereomicroscope (Olympus). Subcellular localization To investigate subcellular localization, the coding regions for BdFAR genes were inserted into the expression vector pA7-GFP by the In-Fusion® HD Cloning Kit (Clontech), for transient expression driven by the CaMV 35S promoter. Resulting C-terminal in-frame fusions with GFP were confirmed by sequencing and then co-transformed into Arabidopsis protoplasts with the ER marker mCherry-HDEL (Nelson et al. 2007) via PEG-mediated transformation as previously described (Bart et al. 2006). The fluorescence signals were detected using a confocal laser-scanning microscope (Leica TCS MP5). Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by the China Postdoctoral Science Foundation [grant No. 2016M602862 to Y.W.]; Fundamental Research Funds for the Central Universities [grant No. 2452016013 to Y.W.]; National Natural Science Foundation of China [grant No. 31271794 to Z.W.]; and Science and Technology Innovation Team Project of Shaanxi Province, China [grant No. 2014KCT-25 to Z.W.]. Acknowledgments We thank Professor Reinhard Jetter for critical reading of the manuscript. Disclosures The authors have no conflicts of interest to declare. References Aarts M.G., Hodge R., Kalantidis K., Florack D., Wilson Z.A., Mulligan B.J., et al.   ( 1997) The Arabidopsis MALE STERILITY 2 protein shares similarity with reductases in elongation/condensation complexes. Plant J.  12: 615– 623. Google Scholar CrossRef Search ADS PubMed  Adamski N.M., Bush M.S., Simmonds J., Turner A.S., Mugford S.G., Jones A., et al.   ( 2013) The Inhibitor of wax 1 locus (Iw1) prevents formation of β- and OH-β-diketones in wheat cuticular waxes and maps to a sub-cM interval on chromosome arm 2BS. Plant J.  74: 989– 1002. Google Scholar CrossRef Search ADS PubMed  Baker E.A. ( 1982) Chemistry and morphology of plant epicuticular waxes. In The Plant Cuticle . Edited by Cutler D., Alvin K.L., Price C.E. pp. 139– 165. Academic Press, London. Barnes J.D., Percy K.E., Paul N.D., Jones P., McLaughlin C.K., Mullineaux P.M., et al.   ( 1996) The influence of UV-B radiation on the physicochemical nature of tobacco (Nicotiana tabacum L.) leaf surfaces. J. Exp. Bot.  47: 99– 109. Google Scholar CrossRef Search ADS   Bart R., Chern M., Park C.J., Bartley L., Ronald P.C. ( 2006) A novel system for gene silencing using siRNAs in rice leaf and stem-derived protoplasts. Plant Methods  2: 13. Google Scholar CrossRef Search ADS PubMed  Barthlott W., Neinhuis C. ( 1997) Purity of the sacred lotus, or escape from contamination in biological surfaces. Planta  202: 1– 8. Google Scholar CrossRef Search ADS   Barthlott W., Neinhuis C., Cutler D., Ditsch F., Meusel I., Theisen I., et al.   ( 1998) Classification and terminology of plant epicuticular waxes. Bot. J. Linn. Soc.  126: 237– 260. Google Scholar CrossRef Search ADS   Beisson F., Li-Beisson Y., Pollard M. ( 2012) Solving the puzzles of cutin and suberin polymer biosynthesis. Curr. Opin. Plant Biol.  15: 329– 337. Google Scholar CrossRef Search ADS PubMed  Bourdenx B., Bernard A., Domergue F., Pascal S., Léger A., Roby D., et al.   ( 2011) Overexpression of Arabidopsis ECERIFERUM1 promotes wax very-long-chain alkane biosynthesis and influences plant response to biotic and abiotic stresses. Plant Physiol.  156: 29– 45. Google Scholar CrossRef Search ADS PubMed  Chen S., Songkumarn P., Liu J., Wang G. ( 2009) A versatile zero background T-vector system for gene cloning and functional genomics. Plant Physiol.  150: 1111– 1121. Google Scholar CrossRef Search ADS PubMed  Chen W., Yu X.H., Zhang K., Shi J., De Oliveira S., Schreiber L., et al.   ( 2011) Male Sterile2 encodes a plastid-localized fatty acyl carrier protein reductase required for pollen exine development in Arabidopsis. Plant Physiol.  157: 842– 853. Google Scholar CrossRef Search ADS PubMed  Cheng J., Russell D.W. ( 2004) Mammalian wax biosynthesis. I. Identification of two fatty acyl-coenzyme A reductases with different substrate specificities and tissue distributions. J. Biol. Chem . 279: 37789– 37797. Google Scholar CrossRef Search ADS PubMed  Clough S.J., Bent A.F. ( 1998) Floral dip: a simplified method for Agrobacterium- mediated transformation of Arabidopsis thaliana. Plant J.  16: 735– 743. Google Scholar CrossRef Search ADS PubMed  Dan Y., Yan H., Munyikwa T., Dong J., Zhang Y., Armstrong C.L. ( 2006) MicroTom: a high-throughput model transformation system for functional genomics. Plant Cell Rep.  25: 432– 441. Google Scholar CrossRef Search ADS PubMed  Doan T.T., Carlsson A.S., Hamberg M., Bülow L., Stymne S., Olsson P. ( 2009) Functional expression of five Arabidopsis fatty acyl-CoA reductase genes in Escherichia coli. J. Plant Physiol.  166: 787– 796. Google Scholar CrossRef Search ADS PubMed  Dobritsa A.A., Shrestha J., Morant M., Pinot F., Matsuno M., Swanson R., et al.   ( 2009) CYP704B1 is a long-chain fatty acid ω-hydroxylase essential for sporopollenin synthesis in pollen of Arabidopsis. Plant Physiol.  151: 574– 589. Google Scholar CrossRef Search ADS PubMed  Domergue F., Vishwanath S.J., Joubès J., Ono J., Lee J.A., Bourdon M., et al.   ( 2010) Three Arabidopsis fatty acyl-coenzyme A reductases, FAR1, FAR4, and FAR5, generate primary fatty alcohols associated with suberin deposition. Plant Physiol.  153: 1539– 1554. Google Scholar CrossRef Search ADS PubMed  Draper J., Mur L.A., Jenkins G., Ghosh-Biswas G.C., Bablak P., Hasterok R., et al.   ( 2001) Brachypodium distachyon. A new model system for functional genomics in grasses. Plant Physiol.  127: 1539– 1555. Google Scholar CrossRef Search ADS PubMed  Eigenbrode S.D., Espelie K.E. ( 1995) Effects of plant epicuticular lipids on insect herbivores. Annu. Rev. Entomol.  40: 171– 194. Google Scholar CrossRef Search ADS   Fillet S., Gibert J., Suárez B., Lara A., Ronchel C., Adrio J.L. ( 2015) Fatty alcohols production by oleaginous yeast. J. Ind. Microbiol. Biotechnol.  42: 1463– 1472. Google Scholar CrossRef Search ADS PubMed  Franke R., Höfer R., Briesen I., Emsermann M., Efremova N., Yephremov A., et al.   ( 2009) The DAISY gene from Arabidopsis encodes a fatty acid elongase condensing enzyme involved in the biosynthesis of aliphatic suberin in roots and the chalaza–micropyle region of seeds. Plant J.  57: 80– 95. Google Scholar CrossRef Search ADS PubMed  Gietz R.D., Woods R.A. ( 2002) Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method. Methods Enzymol . 350: 87– 96. Google Scholar CrossRef Search ADS PubMed  Greer S., Wen M., Bird D., Wu X., Samuels L., Kunst L., et al.   ( 2007) The cytochrome P450 enzyme CYP96A15 is the midchain alkane hydroxylase responsible for formation of secondary alcohols and ketones in stem cuticular wax of Arabidopsis. Plant Physiol.  145: 653– 667. Google Scholar CrossRef Search ADS PubMed  Hall T.A. ( 1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser . 41: 95– 98. Haslam T.M., Mañas-Fernández A., Zhao L.F., Kunst L. ( 2012) Arabidopsis ECERIFERUM2 is a component of the fatty acid elongation machinery required for fatty acid extension to exceptional lengths. Plant Physiol.  160: 1164– 1174. Google Scholar CrossRef Search ADS PubMed  Hegebarth D., Buschhaus C., Joubès J., Thoraval D., Bird D., Jetter R. ( 2017) Arabidopsis ketoacyl-CoA synthase 16 (KCS16) forms C36/C38 acyl precursors for leaf trichome and pavement surface wax. Plant Cell Environ.  40: 1761– 1776. Google Scholar CrossRef Search ADS PubMed  Hofvander P., Doan T.T., Hamberg M. ( 2011) A prokaryotic acyl-CoA reductase performing reduction of fatty acyl-CoA to fatty alcohol. FEBS Lett.  585, 3538– 3543. Google Scholar CrossRef Search ADS PubMed  Hooker T.S., Millar A.A., Kunst L. ( 2002) Significance of the expression of the CER6 condensing enzyme for cuticular wax production in Arabidopsis. Plant Physiol.  129: 1568– 1580. Google Scholar CrossRef Search ADS PubMed  International Brachypodium Initiative. ( 2010) Genome sequencing and analysis of the model grass Brachypodium distachyon. Nature  463: 763– 768. CrossRef Search ADS PubMed  Jefferson R.A., Kavanagh T.A., Bevan M.W. ( 1987) GUS fusions: β-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J.  6: 3901– 3907. Google Scholar PubMed  Jeffree C.E. ( 2007) The fine structure of the plant cuticle. In Annual Plant Reviews 23: Biology of the Plant Cuticle . Edited by Riederer M., Müller C. pp. 11– 125. Blackwell, Oxford. Jetter R., Kunst L., Samuels A.L. ( 2007) Composition of plant cuticular waxes. In Annual Plant Reviews 23: Biology of the Plant Cuticle . Edited by Riederer M., Müller C. pp. 145– 181. Blackwell, Oxford. Joubès J., Raffaele S., Bourdenx B., Garcia C., Laroche-Traineau J., Moreau P., et al.   ( 2008) The VLCFA elongase gene family in Arabidopsis thaliana: phylogenetic analysis, 3D modelling and expression profiling. Plant Mol. Biol.  67: 547– 566. Google Scholar CrossRef Search ADS PubMed  Kim J., Jung J.H., Lee S.B., Go Y.S., Kim H.J., Cahoon R., et al.   ( 2013) Arabidopsis 3-ketoacyl-coenzyme a synthase9 is involved in the synthesis of tetracosanoic acids as precursors of cuticular waxes, suberins, sphingolipids, and phospholipids. Plant Physiol.  162: 567– 580. Google Scholar CrossRef Search ADS PubMed  Kim K.S., Park S.H., Jenks M.A. ( 2007a) Changes in leaf cuticular waxes of sesame (Sesamum indicum L.) plants exposed to water deficit. J. Plant Physiol . 164: 1134– 1143. Google Scholar CrossRef Search ADS   Kim K.S., Park S.H., Kim D.K., Jenks M.A. ( 2007b) Influence of water deficit on leaf cuticular waxes of soybean (Glycine max [L.] Merr.). Int. J. Plant Sci . 168: 307– 316. Google Scholar CrossRef Search ADS   Koch K., Barthlott W., Koch S., Hommes A., Wandelt K., Mamdouh W., et al.   ( 2006) Structural analysis of wheat wax (Triticum aestivum, c.v. ‘Naturastar’ L.): from the molecular level to three dimensional crystals. Planta  223: 258– 270. Google Scholar CrossRef Search ADS PubMed  Kosma D.K., Bourdenx B., Bernard A., Parsons E.P., Lü S., Joubès J., et al.   ( 2009) The impact of water deficiency on leaf cuticle lipids of Arabidopsis. Plant Physiol.  151: 1918– 1929. Google Scholar CrossRef Search ADS PubMed  Kosma D.K., Jenks M.A. ( 2007) Eco-physiological and molecular-genetic determinants of plant cuticle function in drought and salt stress tolerance. In Advances in Molecular Breeding Toward Drought and Salt Tolerant Crops . Edited by Jenks M.A., Hasegawa P.M., Jain S.M. pp. 91– 120. Springer, Dordrecht, The Netherlands. Google Scholar CrossRef Search ADS   Kunst L., Samuels A.L. ( 2003) Biosynthesis and secretion of plant cuticular wax. Prog. Lipid Res.  42: 51– 80. Google Scholar CrossRef Search ADS PubMed  Lee S.B., Jung S.J., Go Y.S., Kim H.U., Kim J.K., Cho H.J., et al.   ( 2009) Two Arabidopsis 3-ketoacyl CoA synthase genes, KCS20 and KCS2/DAISY, are functionally redundant in cuticular wax and root suberin biosynthesis, but differentially controlled by osmotic stress. Plant J.  60: 462– 475. Google Scholar CrossRef Search ADS PubMed  Li F., Wu X., Lam P., Bird D., Zheng H., Samuels L., et al.   ( 2008) Identification of the wax ester synthase/acyl-coenzyme A:diacylglycerol acyltransferase WSD1 required for stem wax ester biosynthesis in Arabidopsis. Plant Physiol.  148: 97– 107. Google Scholar CrossRef Search ADS PubMed  Luna Á. ( 2014) Biosynthesis and accumulation of very-long-chain alkylresorcinols in cuticular waxes of Secale cereale and Brachypodium distachyon. Thesis. University of British Columbia. Mao B., Cheng Z., Lei C., Xu F., Gao S., Ren Y., et al.   ( 2012) Wax crystal-sparse leaf2, a rice homologue of WAX2/GL1, is involved in synthesis of leaf cuticular wax. Planta  235: 39– 52. Google Scholar CrossRef Search ADS PubMed  Metz J.G., Pollard M.R., Anderson L., Hayes T.R., Lassner M.W. ( 2000) Purification of a jojoba embryo fatty acyl-coenzyme A reductase and expression of its cDNA in high erucic acid rapeseed. Plant Physiol.  122: 635– 644. Google Scholar CrossRef Search ADS PubMed  Moto K., Yoshiga T., Yamamoto M., Takahashi S., Okano K., Ando T., et al.   ( 2003) Pheromone gland-specific fatty-acyl reductase of the silkmoth, Bombyx mori. Proc. Natl. Acad. Sci. USA  100: 9156– 9161. Google Scholar CrossRef Search ADS   Nelson B.K., Cai X., Nebenfuhr A. ( 2007) A multicolored set of in vivo organelle markers for co-localization studies in Arabidopsis and other plants. Plant J.  51: 1126– 1136. Google Scholar CrossRef Search ADS PubMed  Panikashvili D., Savaldi-Goldstein S., Mandel T., Yifhar T., Franke R.B., Höfer R., et al.   ( 2007) The Arabidopsis DESPERADO/AtWBC11 transporter is required for cutin and wax secretion. Plant Physiol.  145: 1345– 1360. Google Scholar CrossRef Search ADS PubMed  Peschel S., Franke R., Schreiber L., Knoche M. ( 2007) Composition of the cuticle of developing sweet cherry fruit. Phytochemistry  68: 1017– 1025. Google Scholar CrossRef Search ADS PubMed  Post-Beittenmiller D. ( 1996) Biochemistry and molecular biology of wax production in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol.  47: 405– 430. Google Scholar CrossRef Search ADS PubMed  Raffaele S., Leger A., Roby D. ( 2009) Very long chain fatty acid and lipid signaling in the response of plants to pathogens. Plant Signal. Behav.  4: 94– 99. Google Scholar CrossRef Search ADS PubMed  Rajangam A.S., Gidda S.K., Craddock C., Mullen R.T., Dyer J.M., Eastmond P.J. ( 2013) Molecular characterization of the fatty alcohol oxidation pathway for wax-ester mobilization in germinated jojoba seeds. Plant Physiol.  161: 72– 80. Google Scholar CrossRef Search ADS PubMed  Rowland O., Zheng H., Hepworth S.R., Lam P., Jetter R., Kunst L. ( 2006) CER4 encodes an alcohol-forming fatty acyl-coenzyme A reductase involved in cuticular wax production in Arabidopsis. Plant Physiol.  142: 866– 877. Google Scholar CrossRef Search ADS PubMed  Samuels L., Kunst L., Jetter R. ( 2008) Sealing plant surfaces: cuticular wax formation by epidermal cells. Annu. Rev. Plant Biol.  59: 683– 707. Google Scholar CrossRef Search ADS PubMed  Schneider L.M., Adamski N.M., Christensen C.E., Stuart D.B., Vautrin S., Hansson M., et al.   ( 2016) The Cer-cqu gene cluster determines three key players in a β-diketone synthase polyketide pathway synthesizing aliphatics in epicuticular waxes. J. Exp. Bot.  67: 2715– 2730. Google Scholar CrossRef Search ADS   Seo P.J., Lee S.B., Suh M.C., Park M.J., Go Y.S., Park C.M. ( 2011) The MYB96 transcription factor regulates cuticular wax biosynthesis under drought conditions in Arabidopsis. Plant Cell  23: 1138– 1152. Google Scholar CrossRef Search ADS PubMed  Shepherd T., Wynne Griffiths D. ( 2006) The effects of stress on plant cuticular waxes. New Phytol.  171: 469– 499. Google Scholar CrossRef Search ADS PubMed  Tamura K., Peterson D., Peterson N., Stecher G., Nei M., Kumar S. ( 2011) MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol.  28: 2731– 2739. Google Scholar CrossRef Search ADS PubMed  Teerawanichpan P., Qiu X. ( 2010) Fatty acyl-CoA reductase and wax synthase from Euglena gracilis in the biosynthesis of medium-chain wax esters. Lipids  45: 263– 273. Google Scholar CrossRef Search ADS PubMed  Thompson J.D., Gibson T.J., Plewniak F., Jeanmougin F., Higgins D.G. ( 1997) The CLUSTAL_X Windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res.  25: 4876– 4882. Google Scholar CrossRef Search ADS PubMed  Vogel J., Hill T. ( 2008) High-efficiency Agrobacterium-mediated transformation of Brachypodium distachyon inbred line Bd21-3. Plant Cell Rep.  7: 471– 478. Google Scholar CrossRef Search ADS   Wang A., Xia Q., Xie W., Dumonceaux T., Zou J., Datla R., et al.   ( 2002) Male gametophyte development in bread wheat (Triticum aestivum L.): molecular, cellular, and biochemical analyses of a sporophytic contribution to pollen wall ontogeny. Plant J.  30: 613– 623. Google Scholar CrossRef Search ADS PubMed  Wang M., Wang Y., Wu H., Xu J., Li T., Hegebarth D., et al.   ( 2016) Three TaFAR genes function in the biosynthesis of primary alcohols and the response to abiotic stresses in Triticum aestivum. Sci. Rep.  6: 25008. Google Scholar CrossRef Search ADS PubMed  Wang W., Liu X., Gai X., Ren J., Liu X., Cai Y., et al.   ( 2015a) Cucumis sativus L. WAX2 plays a pivotal role in wax biosynthesis, influencing pollen fertility and plant biotic and abiotic stress responses. Plant Cell Physiol . 56: 1339– 1354. Google Scholar CrossRef Search ADS   Wang W., Wei H., Knoshaug E., Van Wychen S., Xu Q., Himmel M.E., et al.   ( 2016) Fatty alcohol production in Lipomyces starkeyi and Yarrowia lipolytica. Biotechnol. Biofuels  9: 227. Google Scholar CrossRef Search ADS PubMed  Wang W., Zhang Y., Xu C., Ren J., Liu X., Black K., et al.   ( 2015b) Cucumber ECERIFERUM1 (CsCER1), which influences the cuticle properties and drought tolerance of cucumber, plays a key role in VLC alkanes biosynthesis. Plant Mol. Biol . 87: 219– 233. Google Scholar CrossRef Search ADS   Wang Y., Wang J., Chai G., Li C., Hu Y., Chen X., et al.   ( 2015a) Developmental changes in composition and morphology of cuticular waxes on leaves and spikes of glossy and glaucous wheat (Triticum aestivum L.). PLoS One  10: e0141239. Google Scholar CrossRef Search ADS   Wang Y., Wang M., Sun Y., Hegebarth D., Li T., Jetter R., et al.   ( 2015b) Molecular characterization of TaFAR1 involved in primary alcohol biosynthesis of cuticular wax in hexaploid wheat. Plant Cell Physiol . 56: 1944– 1961. Google Scholar CrossRef Search ADS   Wang Y., Wang M., Sun Y., Wang Y., Li T., Chai G., et al.   ( 2015c) FAR5, a fatty acyl-coenzyme A reductase, is involved in primary alcohol biosynthesis of the leaf blade cuticular wax in wheat (Triticum aestivum L.). J. Exp. Bot . 66: 1165– 1178. Google Scholar CrossRef Search ADS   Wang Z., Guhling O., Yao R., Li F., Yeats T.H., Rose J.K., et al.   ( 2011) Two oxidosqualene cyclases responsible for biosynthesis of tomato fruit cuticular triterpenoids. Plant Physiol.  155: 540– 552. Google Scholar CrossRef Search ADS PubMed  Willis R.M., Wahlen B.D., Seefeldt L.C., Barney B.M. ( 2011) Characterization of a fatty acyl-CoA reductase from Marinobacter aquaeolei VT8: a bacterial enzyme catalyzing the reduction of fatty acyl-CoA to fatty alcohol. Biochemistry  50: 10550– 10558. Google Scholar CrossRef Search ADS PubMed  Yeats T.H., Rose J.K. ( 2013) The formation and function of plant cuticles. Plant Physiol.  163: 5– 20. Google Scholar CrossRef Search ADS PubMed  Zhang J., Broeckling C.D., Blancaflor E.B., Sledge M.K., Sumner L.W., Wang Z. ( 2005) Overexpression of WXP1, a putative Medicago truncatula AP2 domain-containing transcription factor gene, increases cuticular wax accumulation and enhances drought tolerance in transgenic alfalfa (Medicago sativa). Plant J.  42: 689– 707. Google Scholar CrossRef Search ADS PubMed  Zhang Z., Wang W., Li W. ( 2013) Genetic interactions underlying the biosynthesis and inhibition of β-diketones in wheat and their impact on glaucousness and cuticle permeability. PLoS One  8: e54129. Google Scholar CrossRef Search ADS PubMed  Zhu X., Xiong L. ( 2013) Putative megaenzyme DWA1 plays essential roles in drought resistance by regulating stress-induced wax deposition in rice. Proc. Natl. Acad. Sci. USA  110: 17790– 17795. Google Scholar CrossRef Search ADS   Abbreviations Abbreviations CaMV Cauliflower mosaic virus ER endoplasmic reticulum FAR fatty acyl-coenzyme A reductase GC-FID gas chromatography with flame ionization detection GC-MS gas chromatography–mass spectroscopy gDNA genomic DNA GFP green fluorescent protein GUS β-glucuronidase PEG polyethylene glycol qRT-PCR quantitative real-time PCR RFP red fluorescent protein SEM scanning electron microscopy VLC very long chain WT wild type Footnotes Footnotes The nucleotide sequences reported in this paper have been submitted to GenBank with accession numbers MF285084, MF285085 and MF285086 © The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Plant and Cell Physiology Oxford University Press

Three Fatty Acyl-Coenzyme A Reductases, BdFAR1, BdFAR2 and BdFAR3, are Involved in Cuticular Wax Primary Alcohol Biosynthesis in Brachypodium distachyon

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Oxford University Press
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© The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com
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0032-0781
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1471-9053
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10.1093/pcp/pcx211
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Abstract

Abstract Plant cuticular wax is a heterogeneous mixture of very long chain fatty acids (VLCFAs) and their derivatives. Primary alcohols are the dominant wax components throughout leaf development of Brachypodium distachyon (Brachypodium). However, the genes involved in primary alcohol biosynthesis have not been investigated and their exact biological function remains unclear in Brachypodium to date. Here, we monitored the leaf wax profile and crystal morphology during Brachypodium leaf morphogenesis, and isolated three Brachypodium fatty acyl-CoA reductase (FAR) genes, named BdFAR1, BdFAR2 and BdFAR3, then analyzed their biochemical activities, substrate specificities, expression patterns, subcellular localization and stress induction. Transgenic expression of BdFAR genes in yeast (Saccharomyces cerevisiae), tomato (Solanum lycopersicum), Arabidopsis (Arabidopsis thaliana) and Brachypodium increased the production of primary alcohols. The three BdFAR genes were preferentially expressed in Brachypodium aerial tissues, consistent with known sites of wax primary alcohol deposition, and localized in the endoplasmic reticulum (ER) in Arabidopsis protoplasts. Finally, expression of the BdFAR genes was induced by drought, cold and ABA treatments, and drought stress significantly increased cuticular wax accumulation in Brachypodium. Taken together, these results indicate that the three BdFAR genes encode active FARs involved in the biosynthesis of Brachypodium wax primary alcohols and respond to abiotic stresses. Introduction The cuticle is a continuous hydrophobic layer that coats most aerial surfaces of terrestrial plants and forms the contact zone between the plant and the environment. The cuticle plays a critical role in limiting non-stomatal water loss (Yeats and Rose 2013) and helps to protect plants against particulate deposits (Barthlott and Neinhuis 1997), UV radiation (Barnes et al. 1996), and bacterial and fungal pathogens (Raffaele et al. 2009). Structurally, the cuticle comprises two major components, cutin and wax. Cutin is a three-dimensional polyester primarily composed of C16 and C18 hydroxy fatty acids, epoxy fatty acids, diacids and glycerol (Beisson et al. 2012), while cuticular wax is a complex mixture commonly including very long chain fatty acids (VLCFAs), primary and secondary alcohols, ketones, aldehydes, alkanes and alkyl esters (Jetter et al. 2007, Samuels et al. 2008). The relative composition of cuticular wax varies greatly among plant species, tissues and different developmental stages of the same organ (Eigenbrode and Espelie 1995, Y. Wang et al. 2015a). The cuticle surfaces of many species and organs are smooth, resulting in a glossy macroscopic appearance, whereas the cuticle surfaces of other species have protruding microscopic wax crystals that give the surface a glaucous appearance (Jeffree 2007). Scanning electron microscopy (SEM) is commonly used to distinguish characteristic wax crystal shapes including platelets, tubules, rodlets and rods (Barthlott et al. 1998), which correlated with certain wax constituents (Jeffree 2007). In diverse plant species and organs, the most important wax compounds are primary alcohols with predominant chain lengths of C26–C30 (Baker 1982). Previous studies have shown that wax alcohols are produced by the acyl reduction pathway in Brassica oleracea L. and involve two separate enzymes with aldehydes as intermediates (Kunst and Samuels 2003). However, subsequent studies have demonstrated that, in Arabidopsis thaliana (Arabidopsis), a single fatty acyl-CoA reductase (FAR) can directly reduce VLC acyl-CoA precursors to primary alcohols (Kunst and Samuels 2003, Samuels et al. 2008). Functional expression of FARs from jojoba (Metz et al. 2000), silkmoth (Bombyx mori) (Moto et al. 2003), mouse (Mus musculus), human (Homo sapiens) (Cheng and Russell 2004), Euglena gracilis (Teerawanichpan and Qiu, 2010) and Marinobacter aquaeolei (Hofvander et al. 2011, Willis et al. 2011, Fillet et al. 2015, W. Wang et al. 2016) in heterologous systems further supported this idea. In recent years, there has been increased interest in FAR proteins that produce primary alcohols. In Arabidopsis, a family of eight putative FAR-like proteins has been identified. AtMS2/FAR2 is involved in the formation of fatty alcohols during pollen exine development (Aarts et al. 1997, Doan et al. 2009, Dobritsa et al. 2009, Chen et al. 2011), while AtCER4/FAR3 catalyzes the synthesis of C24–C30 primary alcohols found in cuticular wax of aerial organs (Rowland et al. 2006). AtFAR1, AtFAR4 and AtFAR5 generate the fatty alcohols found in root, seed coat and wound-induced leaf tissue (Domergue et al. 2010). In wheat, the anther-specific protein TAA1a synthesizes fatty alcohols (Wang et al. 2002). TaFAR1 and TaFAR5 were shown to be involved in cuticular wax primary alcohol biosynthesis in wheat anther and leaf cuticle (Y. Wang et al. 2015b, Y. Wang et al. 2015c). In addition, heterologous expression of the other three wheat TaFARs, TaFAR2, TaFAR3 and TaFAR4, in yeast (Saccharomyces cerevisiae) led to the accumulation of C18:0, C28:0 and C24:0 primary alcohols, respectively (M. Wang et al. 2016). Brachypodium distachyon (Brachypodium) has emerged as an excellent model system to facilitate biological investigations in grasses because of its small genome size, large collection of germplasm resources, short life cycle and simple growing conditions (Draper et al. 2001). The recent release of the genome sequence of the Bd21 accession further accelerates research in this model system (International Brachypodium Initiative 2010). Compared with C28 primary alcohol as the prevalent wax compound in wheat leaf blades (Y. Wang et al. 2015a, Y. Wang et al. 2015b, Y. Wang et al. 2015c), cuticular wax of six Brachypodium organs, i.e. cotyledons, spikes, leaf blades, leaf sheaths, stems and internal stems, was dominated by C26 primary alcohol (Luna 2014). Consequently, the identification of new C26 primary alcohol biosynthetic genes will further enhance our understanding of the substrate range of plant FARs, and this may provide a useful clue for enhancing the C26 primary alcohol production of wheat leaf blades by expressing C26 primary alcohol biosynthetic genes of Brachypodium in future wheat breeding programs. It will be very interesting to determine whether transgenic wheat lines containing high amounts of C26 and C28 primary alcohols could enhance drought tolerance compared with common wheat. In this study, we aimed to monitor the leaf wax profile and surface wax crystals of Brachypodium during leaf development, and to test whether three putative FAR genes, named BdFAR1, BdFAR2 and BdFAR3, are involved in the biosynthesis of cuticular wax primary alcohols for the Brachypodium cuticle. Results Kinetic changes in cuticular wax profiles of developing Brachypodium leaves Total wax was extracted from Brachypodium Bd21 leaves at five growth stages (20, 40, 60, 80 and 100 d).The total wax load increased from 3.77 μg cm−2 at 20 d to 4.85 μg cm−2 at 40 d, and then decreased steadily to 3.13, 2.37 and 1.05 μg cm−2 at 60, 80 and 100 d, respectively (Fig. 1A; Supplementary Table S1). Throughout leaf development, the wax mixture was dominated by primary alcohols (82.40–89.00%), first increasing to 4.32 μg cm−2 at day 40, and then continuously decreasing to 0.87 μg cm−2 at day 100 (Fig. 1A), suggesting that wax biosynthesis of leaves reached a peak at 40 d, especially primary alcohol biosynthesis. They were accompanied by alkanes (7.27–9.99%) and aldehydes (1.78–4.42%), which also increased from 20 to 40 d and then decreased between 60 and 100 d (Fig. 1A). In contrast, fatty acids were present in small amounts (0.94–3.19%) that decreased throughout the entire sampling period (Fig. 1A; Supplementary Table S1). Fig. 1 View largeDownload slide Cuticular wax accumulation and chain length distribution of the individual wax constituents on Bd21 leaves. (A) Developmental changes in the individual wax components and total load. (B) Developmental changes in chain length of the individual wax constituents. Numbers along the x-axis in (B) refer to the total carbon numbers of the compounds. Five representative developmental stages (20, 40, 60, 80 and 100 d) were investigated, and the amounts of cuticular wax are expressed as μg cm−2 of the leaf blade surface area. Each value is the average from three separate samples, and error bars indicate the SD. Fig. 1 View largeDownload slide Cuticular wax accumulation and chain length distribution of the individual wax constituents on Bd21 leaves. (A) Developmental changes in the individual wax components and total load. (B) Developmental changes in chain length of the individual wax constituents. Numbers along the x-axis in (B) refer to the total carbon numbers of the compounds. Five representative developmental stages (20, 40, 60, 80 and 100 d) were investigated, and the amounts of cuticular wax are expressed as μg cm−2 of the leaf blade surface area. Each value is the average from three separate samples, and error bars indicate the SD. The primary alcohols had predominantly even-numbered carbon chain lengths between C22 and C32, peaking at C26 throughout the growth period. Alkanes were found with mainly odd-numbered carbon chain lengths ranging from C25 to C33, with C29 alkane dominating between 20 and 40 d, and C29 and C31 alkanes were the major homologs from 60 to 100 d (Fig. 1B). The major aldehydes detected had C22 and C24 chains, while C26 and C28 aldehydes were present only in trace amounts that could not be quantified. Finally, the major fatty acid had a chain length of C16, accompanied by substantial amounts of C18 fatty acid (Fig. 1B) and traces of C20 and C22 fatty acids. Neither the aldehyde nor the fatty acid chain length profiles changed during leaf ontogenesis. Kinetic changes in wax crystal morphology of developing Brachypodium leaves To visualize the dynamic development of the epicuticular wax crystals on Brachypodium leaf blades, their adaxial and abaxial sides were investigated by SEM at the five growth stages as described above. At 20 d, both leaf sides were covered with very similar platelet-shaped wax crystals that were standing upright, with widely varying angles between them (Supplementary Fig. S1). The size of the wax platelets was typically 0.2–0.4 μm long and 0.1–0.2 μm high (Supplementary Fig. S1). At 40 d, platelet-shaped structures on both the adaxial and abaxial sides had increased lengths of 0.4–0.8 μm and heights of 0.2–0.4 μm, and were arranged in denser networks, suggesting increased numbers of crystals per unit area relative to 20 d. At 60 d, the crystal shapes and arrangements were similar to those at 40 d (Supplementary Fig. S1). During further leaf development, the networks of crystals on both leaf surfaces were gradually thinned, reflecting a decrease in the number of wax crystals compared with leaves at 60 d. Overall, the initial increase and final decrease of wax crystal numbers paralleled similar changes in the total wax load. Isolation and structure analysis of the Brachypodium BdFAR1, BdFAR2, and BdFAR3 genes Based on the finding that Brachypodium cuticular wax was dominated by primary alcohols throughout leaf development (Supplementary Table S1), we hypothesized that homologs of the Arabidopsis CER4 and wheat TaFARs may be important for Brachypodium cuticle formation. To identify FAR-like genes, the Brachypodium genome database was queried with the deduced amino acid sequence of wheat TaFAR1 (GenBank accession No. KF926683) using BLAST search programs. This revealed a group of six Brachypodium FAR-like sequences, of which the three most highly related to TaFAR1 were designated as BdFAR1, BdFAR2 and BdFAR3 (GenBank accession Nos. MF285084, MF285085 and MF285086, respectively). To isolate the BdFAR1, BdFAR2 and BdFAR3 coding regions, reverse transcription–PCR (RT–PCR) was performed using cDNA from Bd21 leaves. BdFAR1, BdFAR2 and BdFAR3 were amplified using the primer pairs BdFAR1-CDS, BdFAR2-CDS and BdFAR3-CDS, respectively, and all primer sequences are listed in Supplementary Table S2. The full-length cDNAs of BdFAR1, BdFAR2 and BdFAR3 are 1,904, 2,012 and 1,856 bp in length, comprising open reading frames (ORFs) of 1,584, 1,533 and 1,494 bp, respectively (Supplementary Fig. S2A–C). The genomic DNA (gDNA) sequence of BdFAR1 spans 3,817 bp with five exons and four introns encoding a 527 amino acid protein. The entire BdFAR2 exon–intron region is 3,931 bp in length and contains eight exons and seven introns encoding a 510 amino acid polypeptide. The gDNA sequence of BdFAR3 spans 4,482 bp, with 10 exons and nine introns encoding a 497 amino acid protein (Supplementary Fig. S2D). Analysis of the BdFAR protein sequences The predicted molecular masses of the BdFAR1, BdFAR2 and BdFAR3 proteins were 58.3, 57.3 and 55.9 kDa, respectively. This predicted result was confirmed by SDS–PAGE analysis of histidine-tagged BdFAR1, BdFAR2 and BdFAR3 proteins (Fig. 2A–C). A predictive protein analysis through the SMART website (http://smart.embl-heidelberg.de/) indicated that the three BdFARs all contained an NAD_binding_4 domain (NADB) at the N-terminus and a male-sterile domain at the C-terminus (Fig. 2D) (Aarts et al. 1997). Previous studies have shown that all plant FARs contained two conserved motifs, an NAD(P)H-binding site motif GXXGXX(G/A) and a classic YXXXK active site motif (Aarts et al. 1997), both of which were also observed in the BdFARs (Fig. 2D). Phylogenic analysis of the 18 plant FARs showed that they can be grouped into three distinct clades (Fig. 2E). The three BdFARs and five wheat TaFARs form the first clade, and seven dicot proteins were grouped into the second clade, including Artemisia annua GFAR1, jojoba ScFAR and Arabidopsis AtFAR1, AtFAR3, AtFAR4, AtFAR5 and AtFAR8. Additionally, the third clade contained rice DPW, Arabidopsis AtFAR2 and AtFAR6. Notably, the FARs in the first and second clades were predicted to be localized in the endoplasmic reticulum (ER), whereas FARs in the third clade were thought to be localized in plastids (Fig. 2E). Overall, the three BdFARs were closely related to the wheat TaFARs involved in primary alcohol biosynthesis for the wheat cuticle, suggesting that the BdFARs play an analogous role in wax alcohol formation in Brachypodium, and thus the biochemical functions of all three BdFARs were further analyzed in the present study. Fig. 2 View large Download slide Sequence analysis of BdFAR1, BdFAR2 and BdFAR3. (A–C) SDS–PAGE of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) proteins. Arrows show the HIS-BdFAR1, HIS-BdFAR2 and HIS-BdFAR3 proteins. (D) Sequence alignment of BdFAR1, BdFAR2 and BdFAR3 proteins. Physicochemically similar residues are shaded in gray. The predicted NAD_binding_4 and active sites are indicated in red boxes. The NAD_binding_4 domain and the male-sterile domain are indicated by black lines and stars under the sequences, respectively. (E) Phylogenetic analysis of plant FARs. This phylogenetic tree was constructed by MEGA 5.0 with the Neighbor–Joining method. The number for each interior branch refers to the percentage of the bootstrap value (1,000 replicates). The GenBank accession numbers of plant FAR genes used in the analysis are summarized in Supplementary Table S3. Fig. 2 View large Download slide Sequence analysis of BdFAR1, BdFAR2 and BdFAR3. (A–C) SDS–PAGE of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) proteins. Arrows show the HIS-BdFAR1, HIS-BdFAR2 and HIS-BdFAR3 proteins. (D) Sequence alignment of BdFAR1, BdFAR2 and BdFAR3 proteins. Physicochemically similar residues are shaded in gray. The predicted NAD_binding_4 and active sites are indicated in red boxes. The NAD_binding_4 domain and the male-sterile domain are indicated by black lines and stars under the sequences, respectively. (E) Phylogenetic analysis of plant FARs. This phylogenetic tree was constructed by MEGA 5.0 with the Neighbor–Joining method. The number for each interior branch refers to the percentage of the bootstrap value (1,000 replicates). The GenBank accession numbers of plant FAR genes used in the analysis are summarized in Supplementary Table S3. Heterologous expression of BdFARs in yeast To determine their biochemical activities and substrate specificities, the BdFAR1–BdFAR3 proteins were first expressed in yeast. To this end, the three coding regions were subcloned into the pYES2 vector for expression under the control of the yeast GAL1 promoter, and the recombinant plasmids were transformed into the yeast mutant strain INVSc1. Gas chromatography–mass spectroscopy (GC-MS) analysis showed that control yeast cells transformed with the empty vector accumulated C16:0, C16:1, C18:0 and C18:1 fatty acids, but no primary alcohols (Fig. 3A). In contrast, the expression of BdFAR1 resulted in the production of C22 alcohol (C22:0-OH), accompanied by small amounts of C24:0-OH (Fig. 3B). Similarly, the expression of BdFAR2 led to the formation of C24:0-OH and C26:0-OH (Fig. 3C), and the expression of BdFAR3 afforded C26:0-OH and minor amounts of C22:0-OH (Fig. 3D). The yeast expression experiments were repeated three times, consistently showing similar results. In summary, these results showed that the three BdFAR proteins have FAR activities to reduce fatty acyl-CoAs to primary alcohols without releasing aldehyde intermediates. The preferred substrate for BdFAR1 is C22:0 fatty acyl-CoA, while that of BdFAR2 and BdFAR3 is C26:0 fatty acyl-CoA, suggesting that both BdFAR2 and BdFAR3 may be involved in the formation of the major Brachypodium leaf wax component, C26 primary alcohol. Fig. 3 View largeDownload slide Heterologous expression of BdFAR1, BdFAR2 and BdFAR3 in yeast. Yeasts were transformed with empty vector control pYES2 (A) or with the pYES2 vector harboring BdFAR1 (B), BdFAR2 (C) or BdFAR3 (D). Major peaks were identified by GC-MS. In the empty vector control, fatty acids (16:1, 16:0, 18:1 and 18:0) but no primary alcohols were detected. In contrast, the yeast strains expressing BdFARs produced novel compounds identified as C22 primary alcohol (1-docosanol; C22:0-OH), C24 primary alcohol (1-tetracosanol; C24:0-OH) and C26 primary alcohol (1-hexacosanol; C26:0-OH). Fig. 3 View largeDownload slide Heterologous expression of BdFAR1, BdFAR2 and BdFAR3 in yeast. Yeasts were transformed with empty vector control pYES2 (A) or with the pYES2 vector harboring BdFAR1 (B), BdFAR2 (C) or BdFAR3 (D). Major peaks were identified by GC-MS. In the empty vector control, fatty acids (16:1, 16:0, 18:1 and 18:0) but no primary alcohols were detected. In contrast, the yeast strains expressing BdFARs produced novel compounds identified as C22 primary alcohol (1-docosanol; C22:0-OH), C24 primary alcohol (1-tetracosanol; C24:0-OH) and C26 primary alcohol (1-hexacosanol; C26:0-OH). Heterologous expression of BdFARs in transgenic tomato leaves and fruits To assess further the biochemical role of BdFAR in planta, we expressed the BdFAR coding regions in tomato (Solanum lycopersicum) cv MicroTom under the control of the Cauliflower mosaic virus (CaMV) 35S promoter via Agrobacterium infiltration (Supplementary Fig. S3A). The transgenic lines carrying the empty vector were used as control. Transgenic plants were screened using hygromycin selection and confirmed by PCR (Supplementary Fig. S3B). All of these transgenic lines displayed similar macroscopic growth phenotypes (Supplementary Fig. S3C). Compared with control plants, all the transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3 had significantly increased amounts of total primary alcohols, whereas the other wax components including n-alkanes, branched alkanes and triterpenoids were not significantly affected (Fig. 4A). In leaves of the transgenic BdFAR1-1 line, the C22:0-OH, C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH contents increased approximately 3.6-, 4.9-, 2.8-, 1.5- and 1.1-fold, respectively, while C32:0-OH was not affected. In the transgenic BdFAR2-2 line, the amounts of C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH increased approximately 6.9-, 4.0-, 2.0- and 1.5-fold, respectively, whereas the amounts of C22:0-OH and C32:0-OH remained nearly the same. Similarly, the transgenic BdFAR3-1 line showed 8.0-, 3.7-, 3.0- and 1.4-fold increases in C24:0-OH, C26:0-OH, C28:0-OH and C30:0-OH, respectively, while C22:0-OH and C32:0-OH were not significantly altered (Fig. 4C). Fig. 4 View largeDownload slide Cuticular wax accumulation in transgenic tomato cv MicroTom. (A, B) Cuticular wax amounts and compound class distribution in the wax of leaves (A) and ripe fruits (B) of T1 transgenic tomato lines. (C, D) Chain length distribution within the primary alcohols in the wax of leaves (C) and ripe fruits (D) of T1 transgenic lines. Control is the empty pCXSN vector plants. Other lines contain the coding regions of the BdFAR genes under the control of the 35S promoter. The data represent the means ± SD of three replicates. Significance is assessed by t-test (*P < 0.05, **P < 0.01). Fig. 4 View largeDownload slide Cuticular wax accumulation in transgenic tomato cv MicroTom. (A, B) Cuticular wax amounts and compound class distribution in the wax of leaves (A) and ripe fruits (B) of T1 transgenic tomato lines. (C, D) Chain length distribution within the primary alcohols in the wax of leaves (C) and ripe fruits (D) of T1 transgenic lines. Control is the empty pCXSN vector plants. Other lines contain the coding regions of the BdFAR genes under the control of the 35S promoter. The data represent the means ± SD of three replicates. Significance is assessed by t-test (*P < 0.05, **P < 0.01). Tomato fruits have wax distinct from that on leaves, with C26:0-OH, C28:0-OH, C30:0-OH and C32:0-OH as major alcohol components, thus enabling further comparative in planta investigations into the effects of BdFAR1–BdFAR3. We found that tomato fruits expressing BdFAR1, BdFAR2 or BdFAR3 had significantly increased amounts of primary alcohols (Fig. 4B), similar to corresponding transgenic leaves. In particular, all the transgenics expressing BdFARs showed increased amounts of C28:0-OH–C32:0-OH, while C34:0-OH remained unchanged relative to the empty vector control. Most interestingly, the lines expressing either BdFAR2 or BdFAR3 had markedly increased amounts of C26:0-OH (Fig. 4D). In addition, we did not observe significant differences in the crystals on leaves and fruits between all transgenic lines and control plants (Supplementary Fig. S4). Taken together, the tomato expression results confirmed that the tested Brachypodium enzymes are active FARs, which accept fairly broad ranges of C24–C32 acyl-CoA substrates. BdFAR2 and BdFAR3 were characterized by relatively strong activity on C26 fatty acyl-CoA, while BdFAR1was distinguished by its ability also to utilize the C22 fatty acyl-CoA substrate. Heterologous expression of BdFARs in Arabidopsis cer4-3 mutant leaves As our yeast and tomato expression results partially diverged, we sought to test further the biochemical characteristics of the BdFAR proteins by heterologous expression in another plant system. To this end, we expressed the BdFAR genes in the Arabidopsis cer4-3 mutant deficient in the formation of C22–C28 primary alcohols (Rowland et al. 2006) and analyzed the wax composition of leaves expressing the BdFAR genes in comparison with empty vector controls. The total amounts of primary alcohols in the BdFAR1 and BdFAR3 transgenic lines were very close to those of the control lines, while the primary alcohol coverages on the BdFAR2 transgenic lines were dramatically increased compared with empty vector control plants (Fig. 5A). All other wax components, including fatty acids, aldehydes, branched alcohols, alkanes, ketone, secondary alcohols and sterols, did not change significantly between the BdFAR transgenic lines and the empty vector control (Fig. 5A). Fig. 5 View largeDownload slide Cuticular wax analysis on leaves of the transgenic Arabidopsis cer4-3 mutant. (A) Distribution of individual compound classes in wax of leaves of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. (B) Chain length distribution within the primary alcohols in the leaf wax of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. Six-week-old Arabidopsis leaves were harvested for cuticular wax analysis. The amounts of wax components are expressed as μg cm−2 leaf surface area. The data represent the means ± SD of three replicates. Asterisks indicate significant differences between the empty vector control and BdFAR1, BdFAR2 or BdFAR3 transgenic lines according to t-test (*P < 0.05; **P < 0.01). Fig. 5 View largeDownload slide Cuticular wax analysis on leaves of the transgenic Arabidopsis cer4-3 mutant. (A) Distribution of individual compound classes in wax of leaves of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. (B) Chain length distribution within the primary alcohols in the leaf wax of an empty vector control line and transgenic lines expressing BdFAR1, BdFAR2 or BdFAR3. Six-week-old Arabidopsis leaves were harvested for cuticular wax analysis. The amounts of wax components are expressed as μg cm−2 leaf surface area. The data represent the means ± SD of three replicates. Asterisks indicate significant differences between the empty vector control and BdFAR1, BdFAR2 or BdFAR3 transgenic lines according to t-test (*P < 0.05; **P < 0.01). Within the primary alcohol fraction in leaf wax of Arabidopsis expressing BdFAR1, the amounts of C22:0-OH, C24:0-OH and C26:0-OH significantly increased, while the amounts of C28:0–C34:0 alcohol did not differ compared with empty vector control plants (Fig. 5B). In the transgenic line expressing BdFAR2, a sharp increase in the content of C26:0-OH was observed along with slightly increased levels of C24:0-OH and C28:0-OH compared with the control (Fig. 5B). Moreover, the expression of BdFAR3 led to increased levels of C22:0-OH, C24:0-OH and C26:0-OH, along with a slight decrease of C32:0-OH relative to control plants, while primary alcohols with other carbon lengths did not show significant changes (Fig. 5B). Together, these results further confirm that BdFARs are active enzymes involved in wax formation in Brachypodium, further suggesting that BdFAR1 and BdFAR2 have preferences for C22 and C26 substrates, respectively. Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves To examine whether BdFAR is involved in the formation of wax alcohols in Brachypodium, we constitutively overexpressed both genes under the control of the CaMV 35S promoter. To this end, the binary vector constructs pCXSN-BdFARs were transformed into Brachypodium (accession Bd21). Unfortunately, our attempt to obtain pCXSN-BdFAR1 transgenic plants was unsuccessful. A total of six independent transgenic BdFAR2 and BdFAR3 lines were obtained. In BdFAR2 and BdFAR3 overexpression lines, the transcript level of BdFAR2 and BdFAR3 significantly increased compared with that in the wild-type (WT) plants (Fig. 6A). The total amounts of alkanes, fatty acids and aldehydes were not affected in the different overexpression lines compared with WT plants, whereas the amounts of primary alcohols were significantly higher in all the overexpression lines (Fig. 6B). In particular, all transgenic lines had levels of C26 alcohol significantly increased relative to the WT, and the amounts of C28 alcohol were also increased in most lines (Fig. 6C). In contrast, the other wax alcohols (C22:0-OH, C24:0-OH, C30:0-OH and C32:0-OH) did not show significant changes between overexpression lines and WT plants (Fig. 6C). Additionally, there were no significant differences in wax crystals on leaves between all transgenic lines and WT plants (Supplementary Fig. S5). Most interestingly, these findings confirmed some but not all of the details in our results from the heterologous expression of the same genes in yeast and other plants. Overall, these results clearly indicated that BdFAR2 and BdFAR3 play a key role in wax alcohol production in Brachypodium, especially the C26 alcohol, which is the major wax component of Brachypodium leaf wax. Fig. 6 View largeDownload slide Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves. (A) qRT-PCR analysis of BdFAR2 and BdFAR3 in WT plants, and BdFAR2- and BdFAR3-overexpressing lines. (B) Distribution of individual compound classes in wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. (C) Chain length distribution within the primary alcohols in the leaf wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. Error bars represent the SD (n =3). The data were statistically analyzed using t-test (*P < 0.05, **P < 0.01). Fig. 6 View largeDownload slide Overexpression of BdFAR2 and BdFAR3 in Brachypodium leaves. (A) qRT-PCR analysis of BdFAR2 and BdFAR3 in WT plants, and BdFAR2- and BdFAR3-overexpressing lines. (B) Distribution of individual compound classes in wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. (C) Chain length distribution within the primary alcohols in the leaf wax of Brachypodium lines overexpressing BdFAR2 or BdFAR3. Error bars represent the SD (n =3). The data were statistically analyzed using t-test (*P < 0.05, **P < 0.01). Temporal and spatial patterns of BdFAR genes in Brachypodium To better understand the functions of BdFAR genes, we examined their spatial and temporal expression patterns in different vegetative and reproductive organs of Bd21. Quantitative real-time PCR (qRT-PCR) showed that the three BdFAR genes were expressed in aerial vegetative and reproductive organs, but not in roots. BdFAR1 was found to be highly expressed in early developing leaves, leaf sheaths, nodes and internodes, modestly expressed in late developing leaves, spikelets and glumes, but not expressed in roots and leaves at 100 d (Fig. 7A). BdFAR2 was mainly expressed in leaf sheaths, nodes, internodes and early developing leaves, and at very low levels in late developing leaves, spikelets and glumes, but not in roots (Fig. 7B). Similarly, the BdFAR3 transcript was detected at high levels in leaves at 40 d, leaf sheaths and internodes, at lower levels in nodes, leaves at 20, 60 and 80 d, spikelets and glumes, but not in roots and leaves at 100 d (Fig. 7C). Notably, the expression of all three BdFAR genes increased greatly between 20 and 40 d, and then gradually decreased during further leaf development, thus paralleling the changes in amounts of primary alcohols on leaf surfaces during leaf development. Fig. 7 View largeDownload slide Temporal and spatial expression patterns of the Brachypodium BdFAR1, BdFAR2 and BdFAR3 genes. (A) Differential expression analysis of BdFAR genes in various organs of Brachypodium by qRT-PCR. LB20, LB40, LB60, LB80 and LB100 represent leaf blades at 20, 40, 60, 80 and 100 d of plant development, respectively. The Brachypodium UBI4 gene was used to normalize gene expression, and error bars represent ± SD of three biological replicates. (B–I) GUS staining analysis of the BdFAR1 (B–E) and the BdFAR2 (F–I) promoter activities in transgenic Brachypodium plants. (B, F) Leaf blades of 50-day-old plants, (C, G) leaf sheaths, (D, H) internodes, (E, I) nodes. Scale bars = 1 mm. Fig. 7 View largeDownload slide Temporal and spatial expression patterns of the Brachypodium BdFAR1, BdFAR2 and BdFAR3 genes. (A) Differential expression analysis of BdFAR genes in various organs of Brachypodium by qRT-PCR. LB20, LB40, LB60, LB80 and LB100 represent leaf blades at 20, 40, 60, 80 and 100 d of plant development, respectively. The Brachypodium UBI4 gene was used to normalize gene expression, and error bars represent ± SD of three biological replicates. (B–I) GUS staining analysis of the BdFAR1 (B–E) and the BdFAR2 (F–I) promoter activities in transgenic Brachypodium plants. (B, F) Leaf blades of 50-day-old plants, (C, G) leaf sheaths, (D, H) internodes, (E, I) nodes. Scale bars = 1 mm. To investigate further the gene expression patterns of BdFAR1 and BdFAR2, genomic promoter sequences 2,264 and 2,285 bp upstream of the start codons, respectively, were fused to the β-glucuronidase (GUS) reporter gene. For both promoters, strong GUS activity was detected in leaf blades (Fig. 7B, F), leaf sheaths (Fig. 7C, G) and nodes (Fig. 7E, I), while only modest GUS activity was observed in internodes (Fig. 7D, H). Unfortunately, our attempt to obtain pBdFAR3:GUS transgenic plants was unsuccessful. Overall, the GUS results are thus consistent with the qRT-PCR data. The expression patterns support the idea that all three BdFARs play a role in the production of primary alcohols for the cuticular wax in Brachypodium. Subcellular localization of BdFAR proteins In silico analyses predicted BdFAR to be localized in the ER (Fig. 2E). To localize the three BdFARs, the corresponding full-length coding sequences without stop codons were fused in-frame to the N-terminus of the green fluorescent protein (GFP) gene for expression under the control of the CaMV 35S promoter, and the resulting constructs were introduced into Arabidopsis leaf protoplasts using polyethylene glycol- (PEG) mediated transformation, and a sequence encoding the ER-specific protein mCherry-HDEL fused with red fluorescent protein (RFP) was co-transformed into the protoplasts (Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016). Both the green BdFAR–GFP and the red HDEL–RFP signals were visualized by confocal microscopy, showing that the GFP fusions of all three BdFARs co-localized completely with the mCherry-HDEL signals (Fig. 8). Consequently, the BdFAR proteins were localized to the ER; thus, the subcellular localization experiments completely confirm the results of the previous in silico analysis (Fig. 2E). Fig. 8 View largeDownload slide Subcellular localization of BdFAR1, BdFAR2 and BdFAR3 in Arabidopsis leaf protoplasts. Each row of five images shows, from left to right, the GFP signal of the BdFAR–GFP fusion construct, the RFP signal of the ER marker mCherry-HDEL, blue fluorescence from the Chl autofluorescence signal, bright-field image and merge of the previous four images. Bars = 5 μm. Fig. 8 View largeDownload slide Subcellular localization of BdFAR1, BdFAR2 and BdFAR3 in Arabidopsis leaf protoplasts. Each row of five images shows, from left to right, the GFP signal of the BdFAR–GFP fusion construct, the RFP signal of the ER marker mCherry-HDEL, blue fluorescence from the Chl autofluorescence signal, bright-field image and merge of the previous four images. Bars = 5 μm. Transcriptional regulation of BdFAR genes under abiotic stress Recent evidence indicated that wax accumulation on the aerial surfaces of diverse plant species could be modulated by water deficiency, cold and ABA treatments (W. Wang et al. 2015a, Y. Wang et al. 2015c, M. Wang et al. 2016). To investigate whether BdFAR expression is regulated by abiotic stresses, we monitored the BdFAR transcript levels in 60-day-old Bd21 seedlings under stress for up to 24 h. When seedlings were subjected to drought, transcript levels of BdFAR1 and BdFAR2 increased approximately 23- and 8-fold, respectively, from the start of the treatment until 24 h (Fig. 9A, B). In contrast, BdFAR3 expression peaked at 2 h, showing a 9-fold increase compared with 0 h (Fig. 9C). Consistent with the drought induction, PEG treatment resulted in approximately 11-, 12- and 9-fold increases in BdFAR1, BdFAR2 and BdFAR3 transcript levels, respectively (Fig. 9A–C). Likewise, cold treatment led to 6-, 4- and 3-fold increases in BdFAR1, BdFAR2 and BdFAR3, respectively (Fig. 9A–C). Based on these observations, we concluded that transcription of the BdFAR genes is positively regulated by abiotic stresses including water deficiency and cold treatments. Next, we examined whether the three BdFAR genes are influenced by the phytohormone ABA. Treatment with 100 μM ABA led to 5-, 23- and 6-fold increases in BdFAR1, BdFAR2 and BdFAR3 transcript abundances, respectively (Fig. 9A–C). From this finding, we concluded that the phytohormone ABA plays an important role in the transcriptional control of BdFAR1–BdFAR3 activation. Fig. 9 View largeDownload slide Expression analysis of BdFAR1, BdFAR2 and BdFAR3 under abiotic stress and ABA treatments. Time course of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) expression after abiotic stress treatment and ABA application. Sixty-day-old soil-grown plants were exposed to various stresses, and relative transcript levels were quantified by qRT-PCR using BdUBI4 as a reference for normalization of the sample amounts. For drought treatment, plants were removed from the soil and allowed to dry under 60% humidity; PEG treatment, 20% (w/v) PEG 6000; Cold treatment, 4°C; ABA treatment, 100 μM ABA. (D) Total wax amounts between well-watered plants (Bd21) and plants after 10 d of water deprivation (Bd21/DR). (E) Cuticular wax composition on leaves of control Bd21 and Bd21/DR plants. Each value represents the mean of three independent measurements. Error bars indicate the SD. Data were statistically analyzed using t-test (*P < 0.05; **P < 0.01). Fig. 9 View largeDownload slide Expression analysis of BdFAR1, BdFAR2 and BdFAR3 under abiotic stress and ABA treatments. Time course of BdFAR1 (A), BdFAR2 (B) and BdFAR3 (C) expression after abiotic stress treatment and ABA application. Sixty-day-old soil-grown plants were exposed to various stresses, and relative transcript levels were quantified by qRT-PCR using BdUBI4 as a reference for normalization of the sample amounts. For drought treatment, plants were removed from the soil and allowed to dry under 60% humidity; PEG treatment, 20% (w/v) PEG 6000; Cold treatment, 4°C; ABA treatment, 100 μM ABA. (D) Total wax amounts between well-watered plants (Bd21) and plants after 10 d of water deprivation (Bd21/DR). (E) Cuticular wax composition on leaves of control Bd21 and Bd21/DR plants. Each value represents the mean of three independent measurements. Error bars indicate the SD. Data were statistically analyzed using t-test (*P < 0.05; **P < 0.01). Studies have shown that the stress-induced expression of wax biosynthesis genes may lead to altered amounts of wax and composition in several plant species (Shepherd and Wynne Griffiths 2006, Panikashvili et al. 2007, Joubès et al. 2008, Kosma et al. 2009). Therefore, we next tested whether the drought effects on BdFAR1–BdFAR3 expression were paralleled by changes in the composition of Brachypodium leaf wax. To this end, we extracted wax from well-irrigated plants (Bd21) and drought-treated plants (Bd21/DR), followed by analysis using GC-MS and gas chromatography with flame ionization detection (GC-FID). After approximately 10 d of drought treatment, leaves of Bd21/DR plants began to wilt (Supplementary Fig. S6). Compared with Bd21, Bd21/DR exhibited a significant increase in the total amount of leaf wax (Fig. 9D), with a 90% increase in fatty acids, an 80% increase in aldehydes, a 60% increase in alkanes and a 60% increase in primary alcohols (Fig. 9E). However, there were no significant shifts in the proportion of these compound classes, or in the chain length distributions within them between Bd21 and Bd21/DR. Altogether, our observations indicate that drought stress induces cuticular wax accumulation in Brachypodium leaves, and that the BdFAR1–BdFAR3 genes play a role in the drought-induced accumulation of cuticular wax, particularly wax alcohols. Discussion Here, we systematically investigated the developmental changes in cuticular wax load, composition and morphology during leaf development in Brachypodium. Our data indicated that primary alcohols were the major components of cuticular wax on Brachypodium leaf blades throughout the entire period of plant growth. Then, we functionally characterized three Brachypodium FAR genes, and showed their involvement in the biosynthesis of cuticular wax primary alcohols. The amounts and crystal densities of cuticular wax vary at different 0developmental stages in Brachypodium In the present study, we observed dynamic changes in wax coverage on Brachypodium leaves, with initially increasing and then continuously decreasing amounts during leaf development (Fig. 1A; Supplementary Table S1). To our knowledge, the present investigation for the first time reports developmental changes of cuticular wax in Brachypodium. Developmental changes in the amount and composition of wax have also been observed in other diverse crops (Peschel et al. 2007, Y. Wang et al. 2015a). The cuticular wax of Brachypodium consisted of the four compound classes, namely primary alcohols, alkanes, aldehydes and fatty acids, and primary alcohols dominated the wax mixture throughout leaf development, as well as primary alcohols being the dominant component in six Brachypodium organs (Luna 2014). We hypothesize that the acyl reduction pathway may play a key role in wax biosynthesis of Brachypodium cuticle. Notably, hexacosanol was the most abundant primary alcohol on Brachypodium leaf surfaces, whereas octacosanol was the predominant primary alcohols on wheat leaves (Adamski et al. 2013, Zhang et al. 2013). In addition, diketone, a key wax component in wheat, barley (Hordeurn vulgare L.) and Arabidopsis (Greer et al. 2007, Y. Wang et al. 2015b, Schneider et al. 2016), has not been detected, suggesting that the glaucous character may be not be present in Brachypodium cuticle. Throughout Brachypodium leaf development, both the adaxial and abaxial leaf surfaces displayed similar platelet-shaped wax crystals (Supplementary Fig. S1). Interestingly, the sizes of the platelets increased, and their arrangements first increased in density then gradually decreased, thus matching the changes in the total amounts of wax. Therefore, our results further support the notion that the formation of platelet crystals is positively correlated with primary alcohols on Brachypodium leaf surfaces. This finding is consistent with previous reports on wax platelets formed by alcohols in other plants (Koch et al. 2006, Zhang et al. 2013, Y. Wang et al. 2015c). Taken together, our findings suggest that differences in surface morphology are driven by dynamic changes in wax alcohol accumulation and underlying quantitative changes in the biosynthesis of the wax alcohols. BdFAR1, BdFAR2 and BdFAR3 are involved in the formation of primary alcohols for the Brachypodium cuticle Here, we isolated three BdFAR genes from Brachypodium. To our knowledge, the BdFAR genes are the first in Brachypodium to be characterized in detail with regard to primary alcohol biosynthesis. Transgenic expression of BdFARs in yeast, tomato, the Arabidopsis cer4-3 mutant and Brachypodium afforded production of primary alcohols, indicating that three BdFAR proteins have FAR activities catalyzing the reduction of fatty acyl-CoAs to primary alcohols. In particular, overexpression of BdFAR2 and BdFAR3 strongly enhanced the amounts of both C26:0 and C28:0 primary alcohol in Brachypodium leaves, suggesting that BdFARs are a promising tool for future studies aiming at enhancing the amounts of plant wax. It is likely that the expression of BdFAR2 and BdFAR3 in wheat may result in high contents of both C26 and C28 primary alcohols and increase drought tolerance. Heterologous expression of yeast indicated that BdFAR1 prefers C22 acyl-CoA as the substrate, whereas BdFAR2 and BdFAR3 preferentially accept C26 acyl-CoA. In fact, BdFAR1 exhibits the highest sequence identity to that of TaFAR1 (75%) and TaFAR5 (74%), which also use C22 acyl-CoA as the preferred substrate in yeast (Y. Wang et al. 2015b, Y. Wang et al. 2015c). Likewise, BdFAR3 shares 84% identity with TaFAR3, which prefers C26 acyl-CoA as the substrate in yeast (M. Wang et al. 2016). However, BdFAR2, TaFAR2 and TaFAR4 are highly similar, but they possess distinct specificities for C26, C24 and C18 acyl-CoAs, respectively (M. Wang et al. 2016). Consequently, it was concluded that sequence similarities of FARs were not correlated with their substrate specificities. The substrate preferences of the BdFAR1, BdFAR2 and BdFAR3 proteins are similar to those of homologous FAR enzymes in other species. For example, Arabidopsis AtFAR1 and AtFAR4 showed preferences for C22 and C20 acyl-CoAs, respectively (Domergue et al. 2010), whereas AtCER4 catalyzes the reduction of C24–C30 acyl-CoAs (Rowland et al. 2006). Jojoba ScFAR uses C20 and C22 acyl-CoAs as substrates (Metz et al. 2000) (Fig. 10). Fig. 10 View largeDownload slide Substrate range of plant FARs involved in synthesis of wax primary alcohols. CER6, KCR1, PAS2, CER10, FAR1, FAR4 and CER4 are wax biosynthetic enzymes from Arabidopsis. ScFAR is the jojoba FAR enzyme involved in the formation of seed storage wax esters. TaFAR1, TaFAR2, TaFAR3, TaFAR4 and TaFAR5 are the FARs involved in wax biosynthesis in wheat. BdFAR1, BdFAR2 and BdFAR3 are the FARs involved in wax biosynthesis in Brachypodium. LACS, long chain acyl-CoA synthetase; FAE, fatty acid elongase. Fig. 10 View largeDownload slide Substrate range of plant FARs involved in synthesis of wax primary alcohols. CER6, KCR1, PAS2, CER10, FAR1, FAR4 and CER4 are wax biosynthetic enzymes from Arabidopsis. ScFAR is the jojoba FAR enzyme involved in the formation of seed storage wax esters. TaFAR1, TaFAR2, TaFAR3, TaFAR4 and TaFAR5 are the FARs involved in wax biosynthesis in wheat. BdFAR1, BdFAR2 and BdFAR3 are the FARs involved in wax biosynthesis in Brachypodium. LACS, long chain acyl-CoA synthetase; FAE, fatty acid elongase. BdFARs are localized to the ER and preferentially expressed in aerial tissues Confocal microscopy observation revealed that BdFAR1, BdFAR2 and BdFAR3 were all localized to the ER (Fig. 8), the subcellular compartment known to harbor all wax biosynthesis reactions (Rowland et al. 2006, Greer et al. 2007, Mao et al. 2012, Rajangam et al. 2013, W. Wang et al. 2015a, Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016, Hegebarth et al. 2017). In Arabidopsis, both WSD1 catalyzing the terminal step of the primary alcohol pathway and the midchain hydroxylase MAH1 catalyzing the final step of the alkane pathway were present in the ER (Greer et al. 2007, Li et al. 2008), leading us to hypothesize that the ER is the core subcellular compartment where almost all cuticular wax components are generated in planta. Further experiments showed that the three investigated BdFAR genes were expressed in all the aerial parts of Brachypodium plants (Fig. 7), consistent with the putative role of these genes in wax production. In particular, the changes in BdFAR expression levels in the course of leaf development were in good accordance with the dynamic changes in the amounts of wax alcohols, further suggesting that the three BdFAR genes are the key enzymes dedicated to wax primary alcohol biosynthesis in Brachypodium. The preferential expression of the BdFARs in aerial organs of Brachypodium is similar to that of many wax biosynthesis genes in other plant species, including Arabidopsis and wheat (Rowland et al. 2006, Greer et al. 2007, Bourdenx et al. 2011, Haslam et al. 2012, Y. Wang et al. 2015c, Y. Wang et al. 2015c, M. Wang et al. 2016). Conversely, it is interesting to note that the three BdFAR genes were barely expressed in roots, suggesting that they are not involved in the formation of suberin or suberin-associated wax. This is in contrast to some of the FARs of Arabidopsis, where AtFAR1, AtFAR4 and AtFAR5 were implicated in suberin biosynthesis (Domergue et al. 2010), along with elongation enzymes encoded, for example, by the ketoacyl-CoA synthase genes KCS2, KCS20 and KCS9 (Franke et al. 2009, Lee et al. 2009, Kim et al. 2013). BdFAR gene expression is regulated by drought, cold stress and ABA treatment Surface wax mixtures play a critical role in protecting plants against biotic and abiotic stresses. One major function of cuticular wax is to prevent non-stomatal water loss, which is especially important for plant survival in water-limited environments (Kosma and Jenks 2007). Accordingly, it has been proposed that during severe water deficit, when stomata are closed, nanoscale diffusion pathways traversing the cuticle become the primary conduit of plant water loss (Kosma et al. 2009). It therefore seems plausible that plants may react to drought stress by modification of their cuticular wax composition, and water deficit may enhance wax synthesis by up-regulating the production of key wax biosynthetic enzymes (Hooker et al. 2002). Our results, showing that expression of the three BdFAR genes was induced under drought stress conditions, clearly support this hypothesis. In addition, BdFAR1 and BdFAR2 were dramatically up-regulated approximately 23- and 8-fold after 24 h of drought stress treatment, and BdFAR3 expression showed a 9-fold increase after 2 h of drought stress treatment, suggesting that induction of expression of BdFAR3 was faster than that of BdFAR1 and BdFAR2 under drought stress. Notably, Brachypodium plants also exhibited a significant increase in the amounts of total leaf wax under drought stress, thus further suggesting that the induction of BdFAR expression indeed leads to changes in the composition of cuticular wax similar to other diverse plant species (Kim et al. 2007a, Kim et al. 2007b, Kosma et al. 2009, Zhu and Xiong, 2013, Y. Wang et al. 2015b). How such changes in cuticle composition may alter its physiological properties and, ultimately, plant performance under drought stress is an important question for future study. Interestingly, ABA treatment also induced BdFAR expression. In Arabidopsis, ABA is known to induce the MYB96 transcription factor which directly binds to conserved sequence motifs in several wax biosynthesis gene promoters and modulates wax biosynthesis in response to drought (Seo et al. 2011). Therefore, it may be speculated that ABA regulation of wax biosynthesis in Brachypodium involves similar transcriptional control mechanisms to those in Arabidopsis, possibly also involving transcription factors such as MYB96. In this context, it should be noted that transcript levels of the BdFAR genes were up-regulated within a few hours under abiotic stresses and ABA treatments, demonstrating that wax production can change rapidly in response to stress. Consistent with our results, the abundances of Arabidopsis CER1 and wheat TaFAR transcripts also increased rapidly under abiotic stress (Bourdenx et al. 2011, Y. Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016). In addition, BdFAR expression was induced by cold treatments, similar to the five wheat TaFAR genes (Wang et al. 2015b, Y. Wang et al. 2015c, M. Wang et al. 2016), cucumber CsWAX2 as well as CsCER1 (W. Wang et al. 2015a, W. Wang et al. 2015b) and Medicago sativa WXP1 (Zhang et al. 2005) which were also up-regulated by cold treatment. These findings are in contrast to Arabidopsis, where cold stress has been found to down-regulate the expression of several components of the FAE complex that are crucial for wax biosynthesis (Joubès et al. 2008). It is likely that the cold stress response varied in severity and effect depending on the plant species. Materials and Methods Plant materials, growth conditions and stress treatments The Brachypodium genotype Bd21 was used for all experiments. Plants were grown in a glasshouse with standard irrigation and fertilization under long-day conditions (18 h of light). Day and night temperatures were 22 and 18°C, respectively. To analyze developmental changes in the wax load, composition and morphology, the first leaves were sampled at 20, 40, 60, 80 and 100 d after germination. Three leaves were used in one extraction and three replicates per stage were tested. Approximately 60 d after germination, Brachypodium plants were subjected to various stress treatments. For drought treatment, plants were air-dried on filter paper and harvested at different time points. For PEG and ABA treatments, plants were transferred to solutions containing 20% (w/v) PEG 6000 and 100 μM ABA, respectively. For cold stress treatment, plants were maintained at 4°C. Cuticular wax extraction and chemical analysis Brachypodium leaves were immersed in 50 ml of chloroform and shaken for 1 min at room temperature. A 20 μg aliquot of n-tetracosane was added as an internal standard. Solvent was evaporated from the resulting extracts under nitrogen, the samples were taken up in chloroform and transferred to a GC autosampler vial, dried again under nitrogen, and derivatized in 100 μl of pyridine and 100 μl of bis-(N, N-trimethylsilyl)-trifluoroacetamide (BSTFA) for 60 min at 70°C. Solvent was again removed under nitrogen and samples were dissolved in 500 μl of chloroform for analyses using GC-MS and GC-FID. The wax composition was determined using a capillary gas chromatograph equipped with an Rxi-5ms column (30 m length, i.d. 0.25 mm, df 0.25 μm; Restek) and a mass spectrometer (GCMS-QP2010, Shimadzu) with helium as the carrier gas. The oven program ran at 50°C for 2 min, increased temperature by 20°C min−1 to 200°C and held for 1 min, increased by 2°C min−1 to 320°C and held for 15 min. Injector and detector temperatures were set at 250°C, and 1 μl of each sample was injected. Identification of wax compounds was based on a comparison of their mass spectra with those of authentic standards and literature data. For the quantification of individual compounds, GC-FID (GC-2010Plus, Shimadzu) with an Rtx-1column (60 m length, i.d. 0.32 mm, df 0.25 μm; Restek) was used under the conditions described above, but with H2 carrier gas. Amounts of all detected compounds were assessed by comparison with peak areas relative to the internal standard, and wax coverage was calculated relative to the extracted surface area calculated by analysis of a digital photograph using Image J software. SEM Fresh leaves were collected from Bd21 plants at 20, 40, 60, 80 and 100 d after germination, air-dried for 10 d in a desiccator at room temperature, and then carefully dissected. Small pieces of samples were mounted on specimen stubs using double-sided copper tape, and sputtered with gold particles at 25 mA for 90 s in a Bal-Tec SCD005 sputter coater (Balzers). The coated surfaces were investigated by SEM (Hitachi S4800) at an accelerating voltage of 10 kV and a working distance of 12 mm. Cloning of BdFAR1, BdFAR2 and BdFAR3 Total RNA was extracted from Bd21 leaves using Trizol Reagent (Invitrogen). Purified RNA was treated with DNase I (Promega) to remove residual DNA, and then was used as the template for first-strand cDNA synthesis using PrimeScript™ reverse transcriptase (TAKARA). The BdFAR cDNAs were amplified with specific primers (Supplementary Table S2) under the following PCR conditions: 95°C for 5 min, 35 cycles of 95°C for 30 s, 58°C for 30 s and 72°C for 2 min, with a final extension at 72°C for 1 min. The amplified products were cloned into the pMD™ 18-T vector (TAKARA) using T4 DNA ligase (TAKARA) and sequenced for verification. The genomic sequences of BdFAR genes were obtained from Brachypodium gDNA. Heterologous expression in yeast The coding sequences of BdFAR genes were amplified from Bd21 cDNA using specific primers (labeled as BdFARx-YS listed in Supplementary Table S2), and the PCR products were cloned into the yeast expression vector pYES2 (Invitrogen) using the In-Fusion® HD Cloning Kit (Clontech). The resulting pYES-BdFARx constructs were transformed into Escherichia coli Top10 cells, and verified by colony PCR and sequencing. Subsequently, the pYES-BdFARx and an empty vector were transformed into the mutant yeast strain INVSc1 according to Gietz and Woods (2002). Transgenic yeast cells were grown on synthetic complete (SC) selection medium without uracil. Three individual cell lines were selected from each transgenic strain. First, yeast cells were inoculated into 20 ml of SC medium containing 2% glucose and grown for 2 d at 30°C, with shaking at 200 r.p.m. Yeast cells were then transferred to 20 ml of induction medium (SC medium containing 2% galactose) for 12 h at 30°C, with shaking at 200 r.p.m. The yeast cells were then transferred to 20 ml of resting medium (0.1 M potassium phosphate containing glucose and hemin) for 24 h at 30°C, with shaking at 200 r.p.m. Then, yeast cells were collected by centrifugation, refluxed for 5 min in 20% KOH/50% ethanol and extracted twice with hexane. Both hexane solutions were combined, and the solvent was removed under a gentle stream of nitrogen. The remaining procedure was consistent with the plant wax analysis described above. Tomato transformation For constitutive tomato expression under the control of the 35S promoter, the coding sequences for BdFAR genes were inserted into a binary vector pCXSN digested with XcmI (NEB) restriction enzyme (Chen et al. 2009). The resulting constructs and an empty vector were individually transformed into Agrobacterium strain GV3101. Cotyledon transformation of tomato cv MicroTom was performed as previously described (Dan et al. 2006). The presence of the transgene was confirmed in the T0 generation by PCR. Transgenic T0 and T1 generation plants were grown in the greenhouse (22°C with 8 h dark/16 h light). Wax analysis of mature leaves and mature red fruits from transgenic plants was performed as previously described (Wang et al. 2011). Arabidopsis transformation The constructs used for tomato transformation (see above) were introduced into the Arabidopsis cer4-3 mutant via Agrobacterium-mediated transformation using the floral dip method (Clough and Bent 1998). The transformed plants were screened on Murashige and Skoog (MS) medium with 25 mg l−1 hygromycin (w/v), and 6-week-old leaves of T1 plants were harvested for cuticular wax analysis. Brachypodium overexpression For BdFAR overexpression under the control of the 35S promoter, the BdFAR2 and BdFAR3 coding regions were amplified and inserted into the XcmI (NEB) site of the pCXSN vector. Then, the resulting constructs were introduced into Agrobacterium strain EHA105, and Brachypodium transformation was performed according to Vogel and Hill (2008). Kanamycin-resistant plants were screened by PCR for transgenes, and selected plantlets were transferred to soil, placed under a plastic wrap for approximately 1 week to acclimate the plants and then grown in the greenhouse. Sequence alignment and phylogenetic analysis The homologous BdFAR proteins were identified in the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov) via BLAST search. Multiple protein sequence alignments were performed with the ClustalW 1.83 program (Thompson et al. 1997), and the aligned sequences were imported into Bioedit for manual editing (http://www.mbio.ncsu.edu/BioEdit/bioedit.html; Hall 1999). A phylogenetic tree was constructed using the Neighbor–Joining (NJ) method by MEGA 5.0 software with the option of pairwise deletion, and 1,000 bootstrap replicates to test inferred phylogeny (Tamura et al. 2011). qRT-PCR For tissue expression analysis, leaf blades from plants at 20, 40, 60, 80 and 100 d, nodes, internodes, leaf sheaths, spikelets, glumes of 9-week-old plants, roots of 2-week-old plants and 60-day-old seedlings treated with various stresses were collected and ground in liquid nitrogen. Total RNA extractions were performed with Trizol Reagent (Invitrogen). After treatment with DNase (Promega), total RNA was used for the first-strand cDNA synthesis using PrimeScript™ reverse transcriptase (TAKARA). qRT-PCR was performed in a 25 μl volume using the SYBR® Premix Ex Taq™ Kit (TAKARA) on a CFX96 real-time PCR detection system (Bio-Rad). BdUBI4 (LOC100832168) was used for normalization of the amount of sample. Each experiment had three biological parallels, each with three technical replicates. Histochemical β-glucuronidase analysis For promoter analysis, the 2,264 and 2,285 bp genomic fragments located upstream of the ATG start codon of BdFAR1 and BdFAR2 were amplified and inserted in front of the GUS gene in pCAMBIA1305 using the In-Fusion® HD Cloning Kit (Clontech). The GUS fusion constructs pBdFAR1:GUS and pBdFAR2:GUS were then transformed into Bd21 plants using Agrobacterium (Vogel and Hill 2008). Histochemical GUS staining was performed as previously described (Jefferson et al. 1987). Transgenic plant samples were incubated in X-gluc buffer at 37°C overnight, and then the staining solution was removed and the samples were cleared of Chl with 70% ethanol. The stained tissues were observed and photographed using a SZX16 stereomicroscope (Olympus). Subcellular localization To investigate subcellular localization, the coding regions for BdFAR genes were inserted into the expression vector pA7-GFP by the In-Fusion® HD Cloning Kit (Clontech), for transient expression driven by the CaMV 35S promoter. Resulting C-terminal in-frame fusions with GFP were confirmed by sequencing and then co-transformed into Arabidopsis protoplasts with the ER marker mCherry-HDEL (Nelson et al. 2007) via PEG-mediated transformation as previously described (Bart et al. 2006). The fluorescence signals were detected using a confocal laser-scanning microscope (Leica TCS MP5). Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by the China Postdoctoral Science Foundation [grant No. 2016M602862 to Y.W.]; Fundamental Research Funds for the Central Universities [grant No. 2452016013 to Y.W.]; National Natural Science Foundation of China [grant No. 31271794 to Z.W.]; and Science and Technology Innovation Team Project of Shaanxi Province, China [grant No. 2014KCT-25 to Z.W.]. Acknowledgments We thank Professor Reinhard Jetter for critical reading of the manuscript. Disclosures The authors have no conflicts of interest to declare. References Aarts M.G., Hodge R., Kalantidis K., Florack D., Wilson Z.A., Mulligan B.J., et al.   ( 1997) The Arabidopsis MALE STERILITY 2 protein shares similarity with reductases in elongation/condensation complexes. Plant J.  12: 615– 623. Google Scholar CrossRef Search ADS PubMed  Adamski N.M., Bush M.S., Simmonds J., Turner A.S., Mugford S.G., Jones A., et al.   ( 2013) The Inhibitor of wax 1 locus (Iw1) prevents formation of β- and OH-β-diketones in wheat cuticular waxes and maps to a sub-cM interval on chromosome arm 2BS. Plant J.  74: 989– 1002. Google Scholar CrossRef Search ADS PubMed  Baker E.A. ( 1982) Chemistry and morphology of plant epicuticular waxes. In The Plant Cuticle . Edited by Cutler D., Alvin K.L., Price C.E. pp. 139– 165. Academic Press, London. Barnes J.D., Percy K.E., Paul N.D., Jones P., McLaughlin C.K., Mullineaux P.M., et al.   ( 1996) The influence of UV-B radiation on the physicochemical nature of tobacco (Nicotiana tabacum L.) leaf surfaces. J. Exp. Bot.  47: 99– 109. Google Scholar CrossRef Search ADS   Bart R., Chern M., Park C.J., Bartley L., Ronald P.C. ( 2006) A novel system for gene silencing using siRNAs in rice leaf and stem-derived protoplasts. Plant Methods  2: 13. Google Scholar CrossRef Search ADS PubMed  Barthlott W., Neinhuis C. ( 1997) Purity of the sacred lotus, or escape from contamination in biological surfaces. Planta  202: 1– 8. Google Scholar CrossRef Search ADS   Barthlott W., Neinhuis C., Cutler D., Ditsch F., Meusel I., Theisen I., et al.   ( 1998) Classification and terminology of plant epicuticular waxes. Bot. J. Linn. Soc.  126: 237– 260. Google Scholar CrossRef Search ADS   Beisson F., Li-Beisson Y., Pollard M. ( 2012) Solving the puzzles of cutin and suberin polymer biosynthesis. Curr. Opin. Plant Biol.  15: 329– 337. Google Scholar CrossRef Search ADS PubMed  Bourdenx B., Bernard A., Domergue F., Pascal S., Léger A., Roby D., et al.   ( 2011) Overexpression of Arabidopsis ECERIFERUM1 promotes wax very-long-chain alkane biosynthesis and influences plant response to biotic and abiotic stresses. Plant Physiol.  156: 29– 45. Google Scholar CrossRef Search ADS PubMed  Chen S., Songkumarn P., Liu J., Wang G. ( 2009) A versatile zero background T-vector system for gene cloning and functional genomics. Plant Physiol.  150: 1111– 1121. Google Scholar CrossRef Search ADS PubMed  Chen W., Yu X.H., Zhang K., Shi J., De Oliveira S., Schreiber L., et al.   ( 2011) Male Sterile2 encodes a plastid-localized fatty acyl carrier protein reductase required for pollen exine development in Arabidopsis. Plant Physiol.  157: 842– 853. Google Scholar CrossRef Search ADS PubMed  Cheng J., Russell D.W. ( 2004) Mammalian wax biosynthesis. I. Identification of two fatty acyl-coenzyme A reductases with different substrate specificities and tissue distributions. J. Biol. Chem . 279: 37789– 37797. Google Scholar CrossRef Search ADS PubMed  Clough S.J., Bent A.F. ( 1998) Floral dip: a simplified method for Agrobacterium- mediated transformation of Arabidopsis thaliana. Plant J.  16: 735– 743. Google Scholar CrossRef Search ADS PubMed  Dan Y., Yan H., Munyikwa T., Dong J., Zhang Y., Armstrong C.L. ( 2006) MicroTom: a high-throughput model transformation system for functional genomics. Plant Cell Rep.  25: 432– 441. Google Scholar CrossRef Search ADS PubMed  Doan T.T., Carlsson A.S., Hamberg M., Bülow L., Stymne S., Olsson P. ( 2009) Functional expression of five Arabidopsis fatty acyl-CoA reductase genes in Escherichia coli. J. Plant Physiol.  166: 787– 796. Google Scholar CrossRef Search ADS PubMed  Dobritsa A.A., Shrestha J., Morant M., Pinot F., Matsuno M., Swanson R., et al.   ( 2009) CYP704B1 is a long-chain fatty acid ω-hydroxylase essential for sporopollenin synthesis in pollen of Arabidopsis. Plant Physiol.  151: 574– 589. Google Scholar CrossRef Search ADS PubMed  Domergue F., Vishwanath S.J., Joubès J., Ono J., Lee J.A., Bourdon M., et al.   ( 2010) Three Arabidopsis fatty acyl-coenzyme A reductases, FAR1, FAR4, and FAR5, generate primary fatty alcohols associated with suberin deposition. Plant Physiol.  153: 1539– 1554. Google Scholar CrossRef Search ADS PubMed  Draper J., Mur L.A., Jenkins G., Ghosh-Biswas G.C., Bablak P., Hasterok R., et al.   ( 2001) Brachypodium distachyon. A new model system for functional genomics in grasses. Plant Physiol.  127: 1539– 1555. Google Scholar CrossRef Search ADS PubMed  Eigenbrode S.D., Espelie K.E. ( 1995) Effects of plant epicuticular lipids on insect herbivores. Annu. Rev. Entomol.  40: 171– 194. Google Scholar CrossRef Search ADS   Fillet S., Gibert J., Suárez B., Lara A., Ronchel C., Adrio J.L. ( 2015) Fatty alcohols production by oleaginous yeast. J. Ind. Microbiol. Biotechnol.  42: 1463– 1472. Google Scholar CrossRef Search ADS PubMed  Franke R., Höfer R., Briesen I., Emsermann M., Efremova N., Yephremov A., et al.   ( 2009) The DAISY gene from Arabidopsis encodes a fatty acid elongase condensing enzyme involved in the biosynthesis of aliphatic suberin in roots and the chalaza–micropyle region of seeds. Plant J.  57: 80– 95. Google Scholar CrossRef Search ADS PubMed  Gietz R.D., Woods R.A. ( 2002) Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method. Methods Enzymol . 350: 87– 96. Google Scholar CrossRef Search ADS PubMed  Greer S., Wen M., Bird D., Wu X., Samuels L., Kunst L., et al.   ( 2007) The cytochrome P450 enzyme CYP96A15 is the midchain alkane hydroxylase responsible for formation of secondary alcohols and ketones in stem cuticular wax of Arabidopsis. Plant Physiol.  145: 653– 667. Google Scholar CrossRef Search ADS PubMed  Hall T.A. ( 1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser . 41: 95– 98. Haslam T.M., Mañas-Fernández A., Zhao L.F., Kunst L. ( 2012) Arabidopsis ECERIFERUM2 is a component of the fatty acid elongation machinery required for fatty acid extension to exceptional lengths. Plant Physiol.  160: 1164– 1174. Google Scholar CrossRef Search ADS PubMed  Hegebarth D., Buschhaus C., Joubès J., Thoraval D., Bird D., Jetter R. ( 2017) Arabidopsis ketoacyl-CoA synthase 16 (KCS16) forms C36/C38 acyl precursors for leaf trichome and pavement surface wax. Plant Cell Environ.  40: 1761– 1776. Google Scholar CrossRef Search ADS PubMed  Hofvander P., Doan T.T., Hamberg M. ( 2011) A prokaryotic acyl-CoA reductase performing reduction of fatty acyl-CoA to fatty alcohol. FEBS Lett.  585, 3538– 3543. Google Scholar CrossRef Search ADS PubMed  Hooker T.S., Millar A.A., Kunst L. ( 2002) Significance of the expression of the CER6 condensing enzyme for cuticular wax production in Arabidopsis. Plant Physiol.  129: 1568– 1580. Google Scholar CrossRef Search ADS PubMed  International Brachypodium Initiative. ( 2010) Genome sequencing and analysis of the model grass Brachypodium distachyon. Nature  463: 763– 768. CrossRef Search ADS PubMed  Jefferson R.A., Kavanagh T.A., Bevan M.W. ( 1987) GUS fusions: β-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J.  6: 3901– 3907. Google Scholar PubMed  Jeffree C.E. ( 2007) The fine structure of the plant cuticle. In Annual Plant Reviews 23: Biology of the Plant Cuticle . Edited by Riederer M., Müller C. pp. 11– 125. Blackwell, Oxford. Jetter R., Kunst L., Samuels A.L. ( 2007) Composition of plant cuticular waxes. In Annual Plant Reviews 23: Biology of the Plant Cuticle . Edited by Riederer M., Müller C. pp. 145– 181. Blackwell, Oxford. Joubès J., Raffaele S., Bourdenx B., Garcia C., Laroche-Traineau J., Moreau P., et al.   ( 2008) The VLCFA elongase gene family in Arabidopsis thaliana: phylogenetic analysis, 3D modelling and expression profiling. Plant Mol. Biol.  67: 547– 566. Google Scholar CrossRef Search ADS PubMed  Kim J., Jung J.H., Lee S.B., Go Y.S., Kim H.J., Cahoon R., et al.   ( 2013) Arabidopsis 3-ketoacyl-coenzyme a synthase9 is involved in the synthesis of tetracosanoic acids as precursors of cuticular waxes, suberins, sphingolipids, and phospholipids. Plant Physiol.  162: 567– 580. Google Scholar CrossRef Search ADS PubMed  Kim K.S., Park S.H., Jenks M.A. ( 2007a) Changes in leaf cuticular waxes of sesame (Sesamum indicum L.) plants exposed to water deficit. J. Plant Physiol . 164: 1134– 1143. Google Scholar CrossRef Search ADS   Kim K.S., Park S.H., Kim D.K., Jenks M.A. ( 2007b) Influence of water deficit on leaf cuticular waxes of soybean (Glycine max [L.] Merr.). Int. J. Plant Sci . 168: 307– 316. Google Scholar CrossRef Search ADS   Koch K., Barthlott W., Koch S., Hommes A., Wandelt K., Mamdouh W., et al.   ( 2006) Structural analysis of wheat wax (Triticum aestivum, c.v. ‘Naturastar’ L.): from the molecular level to three dimensional crystals. Planta  223: 258– 270. Google Scholar CrossRef Search ADS PubMed  Kosma D.K., Bourdenx B., Bernard A., Parsons E.P., Lü S., Joubès J., et al.   ( 2009) The impact of water deficiency on leaf cuticle lipids of Arabidopsis. Plant Physiol.  151: 1918– 1929. Google Scholar CrossRef Search ADS PubMed  Kosma D.K., Jenks M.A. ( 2007) Eco-physiological and molecular-genetic determinants of plant cuticle function in drought and salt stress tolerance. In Advances in Molecular Breeding Toward Drought and Salt Tolerant Crops . Edited by Jenks M.A., Hasegawa P.M., Jain S.M. pp. 91– 120. Springer, Dordrecht, The Netherlands. Google Scholar CrossRef Search ADS   Kunst L., Samuels A.L. ( 2003) Biosynthesis and secretion of plant cuticular wax. Prog. Lipid Res.  42: 51– 80. Google Scholar CrossRef Search ADS PubMed  Lee S.B., Jung S.J., Go Y.S., Kim H.U., Kim J.K., Cho H.J., et al.   ( 2009) Two Arabidopsis 3-ketoacyl CoA synthase genes, KCS20 and KCS2/DAISY, are functionally redundant in cuticular wax and root suberin biosynthesis, but differentially controlled by osmotic stress. Plant J.  60: 462– 475. Google Scholar CrossRef Search ADS PubMed  Li F., Wu X., Lam P., Bird D., Zheng H., Samuels L., et al.   ( 2008) Identification of the wax ester synthase/acyl-coenzyme A:diacylglycerol acyltransferase WSD1 required for stem wax ester biosynthesis in Arabidopsis. Plant Physiol.  148: 97– 107. Google Scholar CrossRef Search ADS PubMed  Luna Á. ( 2014) Biosynthesis and accumulation of very-long-chain alkylresorcinols in cuticular waxes of Secale cereale and Brachypodium distachyon. Thesis. University of British Columbia. Mao B., Cheng Z., Lei C., Xu F., Gao S., Ren Y., et al.   ( 2012) Wax crystal-sparse leaf2, a rice homologue of WAX2/GL1, is involved in synthesis of leaf cuticular wax. Planta  235: 39– 52. Google Scholar CrossRef Search ADS PubMed  Metz J.G., Pollard M.R., Anderson L., Hayes T.R., Lassner M.W. ( 2000) Purification of a jojoba embryo fatty acyl-coenzyme A reductase and expression of its cDNA in high erucic acid rapeseed. Plant Physiol.  122: 635– 644. Google Scholar CrossRef Search ADS PubMed  Moto K., Yoshiga T., Yamamoto M., Takahashi S., Okano K., Ando T., et al.   ( 2003) Pheromone gland-specific fatty-acyl reductase of the silkmoth, Bombyx mori. Proc. Natl. Acad. Sci. USA  100: 9156– 9161. Google Scholar CrossRef Search ADS   Nelson B.K., Cai X., Nebenfuhr A. ( 2007) A multicolored set of in vivo organelle markers for co-localization studies in Arabidopsis and other plants. Plant J.  51: 1126– 1136. Google Scholar CrossRef Search ADS PubMed  Panikashvili D., Savaldi-Goldstein S., Mandel T., Yifhar T., Franke R.B., Höfer R., et al.   ( 2007) The Arabidopsis DESPERADO/AtWBC11 transporter is required for cutin and wax secretion. Plant Physiol.  145: 1345– 1360. Google Scholar CrossRef Search ADS PubMed  Peschel S., Franke R., Schreiber L., Knoche M. ( 2007) Composition of the cuticle of developing sweet cherry fruit. Phytochemistry  68: 1017– 1025. Google Scholar CrossRef Search ADS PubMed  Post-Beittenmiller D. ( 1996) Biochemistry and molecular biology of wax production in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol.  47: 405– 430. Google Scholar CrossRef Search ADS PubMed  Raffaele S., Leger A., Roby D. ( 2009) Very long chain fatty acid and lipid signaling in the response of plants to pathogens. Plant Signal. Behav.  4: 94– 99. Google Scholar CrossRef Search ADS PubMed  Rajangam A.S., Gidda S.K., Craddock C., Mullen R.T., Dyer J.M., Eastmond P.J. ( 2013) Molecular characterization of the fatty alcohol oxidation pathway for wax-ester mobilization in germinated jojoba seeds. Plant Physiol.  161: 72– 80. Google Scholar CrossRef Search ADS PubMed  Rowland O., Zheng H., Hepworth S.R., Lam P., Jetter R., Kunst L. ( 2006) CER4 encodes an alcohol-forming fatty acyl-coenzyme A reductase involved in cuticular wax production in Arabidopsis. Plant Physiol.  142: 866– 877. Google Scholar CrossRef Search ADS PubMed  Samuels L., Kunst L., Jetter R. ( 2008) Sealing plant surfaces: cuticular wax formation by epidermal cells. Annu. Rev. Plant Biol.  59: 683– 707. Google Scholar CrossRef Search ADS PubMed  Schneider L.M., Adamski N.M., Christensen C.E., Stuart D.B., Vautrin S., Hansson M., et al.   ( 2016) The Cer-cqu gene cluster determines three key players in a β-diketone synthase polyketide pathway synthesizing aliphatics in epicuticular waxes. J. Exp. Bot.  67: 2715– 2730. Google Scholar CrossRef Search ADS   Seo P.J., Lee S.B., Suh M.C., Park M.J., Go Y.S., Park C.M. ( 2011) The MYB96 transcription factor regulates cuticular wax biosynthesis under drought conditions in Arabidopsis. Plant Cell  23: 1138– 1152. Google Scholar CrossRef Search ADS PubMed  Shepherd T., Wynne Griffiths D. ( 2006) The effects of stress on plant cuticular waxes. New Phytol.  171: 469– 499. Google Scholar CrossRef Search ADS PubMed  Tamura K., Peterson D., Peterson N., Stecher G., Nei M., Kumar S. ( 2011) MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol.  28: 2731– 2739. Google Scholar CrossRef Search ADS PubMed  Teerawanichpan P., Qiu X. ( 2010) Fatty acyl-CoA reductase and wax synthase from Euglena gracilis in the biosynthesis of medium-chain wax esters. Lipids  45: 263– 273. Google Scholar CrossRef Search ADS PubMed  Thompson J.D., Gibson T.J., Plewniak F., Jeanmougin F., Higgins D.G. ( 1997) The CLUSTAL_X Windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res.  25: 4876– 4882. Google Scholar CrossRef Search ADS PubMed  Vogel J., Hill T. ( 2008) High-efficiency Agrobacterium-mediated transformation of Brachypodium distachyon inbred line Bd21-3. Plant Cell Rep.  7: 471– 478. Google Scholar CrossRef Search ADS   Wang A., Xia Q., Xie W., Dumonceaux T., Zou J., Datla R., et al.   ( 2002) Male gametophyte development in bread wheat (Triticum aestivum L.): molecular, cellular, and biochemical analyses of a sporophytic contribution to pollen wall ontogeny. Plant J.  30: 613– 623. Google Scholar CrossRef Search ADS PubMed  Wang M., Wang Y., Wu H., Xu J., Li T., Hegebarth D., et al.   ( 2016) Three TaFAR genes function in the biosynthesis of primary alcohols and the response to abiotic stresses in Triticum aestivum. Sci. Rep.  6: 25008. Google Scholar CrossRef Search ADS PubMed  Wang W., Liu X., Gai X., Ren J., Liu X., Cai Y., et al.   ( 2015a) Cucumis sativus L. WAX2 plays a pivotal role in wax biosynthesis, influencing pollen fertility and plant biotic and abiotic stress responses. Plant Cell Physiol . 56: 1339– 1354. Google Scholar CrossRef Search ADS   Wang W., Wei H., Knoshaug E., Van Wychen S., Xu Q., Himmel M.E., et al.   ( 2016) Fatty alcohol production in Lipomyces starkeyi and Yarrowia lipolytica. Biotechnol. Biofuels  9: 227. Google Scholar CrossRef Search ADS PubMed  Wang W., Zhang Y., Xu C., Ren J., Liu X., Black K., et al.   ( 2015b) Cucumber ECERIFERUM1 (CsCER1), which influences the cuticle properties and drought tolerance of cucumber, plays a key role in VLC alkanes biosynthesis. Plant Mol. Biol . 87: 219– 233. Google Scholar CrossRef Search ADS   Wang Y., Wang J., Chai G., Li C., Hu Y., Chen X., et al.   ( 2015a) Developmental changes in composition and morphology of cuticular waxes on leaves and spikes of glossy and glaucous wheat (Triticum aestivum L.). PLoS One  10: e0141239. Google Scholar CrossRef Search ADS   Wang Y., Wang M., Sun Y., Hegebarth D., Li T., Jetter R., et al.   ( 2015b) Molecular characterization of TaFAR1 involved in primary alcohol biosynthesis of cuticular wax in hexaploid wheat. Plant Cell Physiol . 56: 1944– 1961. Google Scholar CrossRef Search ADS   Wang Y., Wang M., Sun Y., Wang Y., Li T., Chai G., et al.   ( 2015c) FAR5, a fatty acyl-coenzyme A reductase, is involved in primary alcohol biosynthesis of the leaf blade cuticular wax in wheat (Triticum aestivum L.). J. Exp. Bot . 66: 1165– 1178. Google Scholar CrossRef Search ADS   Wang Z., Guhling O., Yao R., Li F., Yeats T.H., Rose J.K., et al.   ( 2011) Two oxidosqualene cyclases responsible for biosynthesis of tomato fruit cuticular triterpenoids. Plant Physiol.  155: 540– 552. Google Scholar CrossRef Search ADS PubMed  Willis R.M., Wahlen B.D., Seefeldt L.C., Barney B.M. ( 2011) Characterization of a fatty acyl-CoA reductase from Marinobacter aquaeolei VT8: a bacterial enzyme catalyzing the reduction of fatty acyl-CoA to fatty alcohol. Biochemistry  50: 10550– 10558. Google Scholar CrossRef Search ADS PubMed  Yeats T.H., Rose J.K. ( 2013) The formation and function of plant cuticles. Plant Physiol.  163: 5– 20. Google Scholar CrossRef Search ADS PubMed  Zhang J., Broeckling C.D., Blancaflor E.B., Sledge M.K., Sumner L.W., Wang Z. ( 2005) Overexpression of WXP1, a putative Medicago truncatula AP2 domain-containing transcription factor gene, increases cuticular wax accumulation and enhances drought tolerance in transgenic alfalfa (Medicago sativa). Plant J.  42: 689– 707. Google Scholar CrossRef Search ADS PubMed  Zhang Z., Wang W., Li W. ( 2013) Genetic interactions underlying the biosynthesis and inhibition of β-diketones in wheat and their impact on glaucousness and cuticle permeability. PLoS One  8: e54129. Google Scholar CrossRef Search ADS PubMed  Zhu X., Xiong L. ( 2013) Putative megaenzyme DWA1 plays essential roles in drought resistance by regulating stress-induced wax deposition in rice. Proc. Natl. Acad. Sci. USA  110: 17790– 17795. Google Scholar CrossRef Search ADS   Abbreviations Abbreviations CaMV Cauliflower mosaic virus ER endoplasmic reticulum FAR fatty acyl-coenzyme A reductase GC-FID gas chromatography with flame ionization detection GC-MS gas chromatography–mass spectroscopy gDNA genomic DNA GFP green fluorescent protein GUS β-glucuronidase PEG polyethylene glycol qRT-PCR quantitative real-time PCR RFP red fluorescent protein SEM scanning electron microscopy VLC very long chain WT wild type Footnotes Footnotes The nucleotide sequences reported in this paper have been submitted to GenBank with accession numbers MF285084, MF285085 and MF285086 © The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com

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Plant and Cell PhysiologyOxford University Press

Published: Mar 1, 2018

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