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Biosynthesis and accumulation of very-long-chain alkylresorcinols in cuticular waxes of Secale cereale… Luna, Álvaro 2014

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 BIOSYNTHESIS AND ACCUMULATION OF VERY-LONG-CHAIN ALKYLRESORCINOLS IN CUTICULAR WAXES OF  Secale cereale AND Brachypodium distachyon  by  Álvaro Luna  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE  in The Faculty of Graduate and Postdoctoral Studies  (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver) July 2014   © Álvaro Luna, 2014 ii  Abstract  5-Alkylresorcinols are a class of phenolic lipids which have been identified in the cuticular waxes of various cereal crops. Due to their antifungal and antibacterial properties, ARs have potential applications as nutraceuticals. They are biosynthesized by type-III polyketide synthases (PKSs). Two candidate PKS genes were previously isolated from the two model grass species Secale cereale and Brachypodium distachyon, and were shown to encode alkylresorcinol synthases (ScARS and BdARS, respectively). Here I report the further characterization of these two enzymes, with the goal to test whether they are involved in the formation of cuticular wax alkylresorcinols. Series of alkylresorcinols were identified and quantified, containing ARs with C19-C27 alkyl chains in S. cereale waxes, and C17-C25 on B. distachyon waxes. In addition, a new series of methyl-branched alkylresorcinols was identified with C19-C25 chains. The accumulation of ARs was monitored in waxes on various organs of etiolated and normal plants, and the product amounts found to correlate with the expression patterns of the putative ARS genes in each species. Subcellular localization using GFP fusions showed that the ARS proteins are associated with ER membranes of epidermal cells, where very-long-chain acyl CoA substrates of ARSs are known to accumulate. Overall, my data indicate that both enzymes are indeed involved in the biosynthesis of grass surface alkylresorcinols.     iii  Preface  The work in Chapter 4 was based on previous research performed by Ruonan Yao, who isolated the two putative genes involved in the biosynthesis of alkylresorcinols in Secale cereale and Brachypodium distachyon. The synthetic alkylresorcinol used in this thesis was performed by former student Xiufen Ji. The synthesis of methyl alkylresorcinol standard was conducted in collaboration with Chen Peng. The subcellular localization experiment of ScARS and BdARS was conducted partially in collaboration with Mariya Skvortsova, who gave technical instructions, and Sandra Keerthisinhe, who manipulated the microscope.   All the wax analyses, identification and quantification were my contribution, as well as the manuscript of this thesis. All the revisions of the current thesis were performed with the help of Dr. Reinhard Jetter.             iv  Table of Contents  Abstract .................................................................................................................................... ii Preface ..................................................................................................................................... iii Table of Contents ................................................................................................................... iv List of Tables ......................................................................................................................... vii List of Figures ....................................................................................................................... viii List of Abbreviations ............................................................................................................ xii Acknowledgements .............................................................................................................. xiv Dedication ………………………………………………………………………………….xv   Chapter 1: Introduction ......................................................................................................... 1 1.1 Plant cuticular waxes .................................................................................................... 1 1.1.1 The plant cuticle ....................................................................................................... 1 1.1.2 Structure and composition of the plant cuticle ........................................................ 1 1.1.3 Wax biosynthesis ..................................................................................................... 3 1.2Alkylresorcinols .............................................................................................................. 4 1.2.1 Occurrence and biological functions of alkylresorcinols ......................................... 4 1.2.2   Biosynthesis of alkylresorcinols ............................................................................ 7 1.2.3 Secale cereale and Brachypodium distachyon as model system ........................... 10 1.2.4 Isolation and partial characterization of BdARS and ScARS ................................ 11 1.2.5   Effects of etiolation on cuticular alkylresorcinols ............................................... 13 1.3 Research questions and objectives ............................................................................ 14  Chapter 2: Materials and methods ...................................................................................... 16 2.1 Plant material and growth conditions ....................................................................... 16 2.1.1 Plant growth conditions ......................................................................................... 16 2.1.2 Plant materials ........................................................................................................ 16 2.2 Wax analysis ................................................................................................................ 17 2.2.1 Wax extraction and derivatization ......................................................................... 17 2.2.2 Chemical analysis by GC-FID and GC-MS ........................................................... 18 2.2.3 Synthesis of C19 methyl alkylresorcinol (CMAR19:0) .......................................... 19 2.3 Genetic analysis ........................................................................................................... 21 2.3.1 RNA isolation and reverse transcription ................................................................ 21 2.3.2 Semi quantitative RT-PCR .................................................................................... 22 v  2.3.3 Agrobacterium-mediated infiltration ..................................................................... 22  Chapter 3: Total wax profiles in various organs of Secale cereale and Brachypodium distachyon ............................................................................................................................... 24 3.1 Introduction ................................................................................................................. 24 3.2 Results .......................................................................................................................... 28 3.2.1 Cuticular wax profiles of various S. cereale organs .............................................. 28 3.2.1.2 Total wax amounts on S. cereale organs ......................................................... 30 3.2.1.3 Compound class distribution in waxes on etiolated and green S. cereale organs ..................................................................................................................................... 31 3.2.1.4 Chain length distributions in waxes from etiolated and green S. cereale organs ..................................................................................................................................... 34 3.2.2 Cuticular wax profile of various Brachypodium distachyon organs. ..................... 37 3.2.2.1 Total wax amount of Brachypodium distachyon organs ................................ 37 3.2.2.2 Compound class distribution in waxes on Brachypodium distachyon organs 38 3.2.2.3 Chain length distribution in waxes from Brachypodium distachyon organs .. 40 3.3 Discussion ..................................................................................................................... 42 3.3.1 Total wax of green and etiolated S. cereale organs ............................................... 42 3.3.2 Compound class and homolog distributions in cuticular waxes from S. cereale organs .............................................................................................................................. 43 3.3.4 Total wax of Brachypodium distachyon organs ..................................................... 48 3.3.5 Homolog distributions in cuticular waxes from Brachypodium distachyon organs ......................................................................................................................................... 48  Chapter 4: Cuticular alkylresorcinol homologs in various organs of Secale cereale and Brachypodium distachyon ..................................................................................................... 50 4.1 Introduction ................................................................................................................. 50 4.2 Results .......................................................................................................................... 54 4.2.1 Identification of cuticular alkylresorcinols and methyl alkylresorcinols ............... 54 4.2.1.1 Identification of alkylresorcinol homologs in waxes from Secale cereale and B. distachyon ............................................................................................................... 55 4.2.1.2 Identification of methyl alkylresorcinols in waxes from S. cereale and B. distachyon ................................................................................................................... 60 4.2.2 Quantification of alkylresorcinols in waxes from various Secale cereale and B. distachyon organs ............................................................................................................ 65 4.2.2.1 Alkylresorcinol chain length distribution in Secale cereale and B. distachyon organs .......................................................................................................................... 65 vi  4.2.2.2 Methyl alkylresorcinol chain length distribution in Secale cereale and B. distachyon organs ........................................................................................................ 67 4.2.2.3 Total cuticular alkylresorcinol amounts .......................................................... 69 4.2.3 Gene expression ..................................................................................................... 70 4.2.4 Subcellular localization. ......................................................................................... 71 4.3 Discussion ..................................................................................................................... 74 4.3.1 Identification of cuticular alkylresorcinols and methyl alkylresorcinols ............... 74 4.3.2 Homolog distributions of cuticular alkylresorcinols in Secale cereale and B. distachyon waxes ............................................................................................................ 79 4.3.3 Total amounts of cuticular alkylresorcinols ........................................................... 81 4.3.4 Gene expression analysis and subcellular localization of ScARS and BdARS ..... 82  Chapter 5: Alkyl ester compositions on green and etiolated organs of Secale cereale ... 84 5.1 Introduction ................................................................................................................. 84 5.2 Results .......................................................................................................................... 88 5.2.1 Alkyl ester composition in waxes from Secale cereale organs ............................. 88 5.2.1.1 Total cuticular alkyl ester amounts ................................................................. 88 5.2.1.2 Alkyl ester chain length distributions on green and etiolated Secale cereale organs .......................................................................................................................... 89 5.2.2 Chain length distributions of acyl and alkyl moieties in Secale cereale wax esters ......................................................................................................................................... 90 5.3 Discussion ..................................................................................................................... 94 5.3.1 Total alkyl ester amounts in waxes from Secale cereale organs ........................... 94 5.3.2 Alkyl ester chain length distribution in green and etiolated Secale cereale organs ......................................................................................................................................... 95 5.3.3 Acyl and alkyl moieties in wax esters from selected green and etiolated Secale cereale organs ................................................................................................................. 96 5.3.4 Chain length distribution of free and esterified alcohol and fatty acids ................ 97  Chapter 6: Conclusions and future directions ................................................................. 100 6.1 The wax biosynthesis on various organs of Secale cereale and Brachypodium distachyon ......................................................................................................................... 100 6.2 Biosynthesis of cuticular alkylresorcinols .................................................................. 103 6.3 Methyl alkylresorcinols in grasses .............................................................................. 105  References ............................................................................................................................ 107  vii  List of Tables Table 4.1 List of alkylresorcinol homologs identified in various S. cereale and B. distachyon organs. .......................................................................................................................... 59 Table 4.2 List of methyl alkylresorcinol homologs identified in various S. cereale and B. distachyon organs. ....................................................................................................... 62 Table 5.1 Compound class composition of waxes from various Secale cereale organs. Relative amounts [%] of compound classes are given as mean values (n = 6). .......... 89 Table 5.2 Relative amounts [%] of acyl moieties in wax esters from various S. cereale organs. .......................................................................................................................... 93                 viii  List of Figures Figure 1.1 Schematic cross-section of the plant epidermis and cuticle (Modified from Jetter et al., 2000). ................................................................................................................... 2 Figure 1.2 Structure of 5-n-alkylresorcinol (n=1,2,3…). ......................................................... 5 Figure 1.3 Chalcone synthase and stilbene synthase mechanism for forming naringenin chalcone and resveratrol using Claisen and aldol condensations, respectively. ............ 8 Figure 1.4 Possible mechanism for the formation of alkylresorcinols through elongation and aldol condensation catalyzed by alkylresorcinol synthase (ARS). ................................ 9 Figure 1.5 Gene structures of previously isolated BdARS from B. distachyon and ScARS from S. cereale. Exons are shown in shaded boxes and introns are shown in lines (Adapted from Yao, 2010). .......................................................................................... 12 Figure 2.1 Synthesis of C21 methyl alkylresorcinol with methylation on the benzylic carbon. ..................................................................................................................................... 20 Figure 3.1 Growth of green and etiolated S. cereale leaf blades and sheaths. Blades and sheaths were distinguished as the parts of the leaf above and below the POE (point of emergence). Growth was monitored by measuring blade and sheath length in situ until day 20 after germination. Data are given as mean values (n=4) ± SD. ....................... 30 Figure 3.2 Total wax loads on various S. cereale organs grown in the light or in the dark. Wax coverages are given as mean values (n = 6) ± SD. ............................................. 31 Figure 3.3 Compound class compositions of wax mixtures from various S. cereale organs. Waxes from green leaf blades, external sheaths and internal sheaths, and from etiolated leaf blades, stems, hypocotyls, cotyledons and sheaths were analyzed. The coverage of each compound class is given as mean value in µg/cm2 ± SD. ............... 33 Figure 3.4 Chain length distributions within compound classes in waxes from green S. cereale organs. Relative amounts of homologs are given as % of the wax fraction from green leaf blades, external and internal sheaths. The percentage of each homolog within the compound class is given as mean value (n = 6) ± SD. ............................... 35 Figure 3.5 Chain length distributions within compound classes in waxes from etiolated S. cereale organs. Relative amounts of homologs are given as % of the fraction within waxes from leaf blades, stems, sheaths, hypocotyls and cotyledons. The percentage of each homolog within the compound class is given as mean value (n = 6) ± SD. ....... 36 Figure 3.6 Total wax loads on various B. distachyon organs. Wax coverages are given as mean values (n = 6) ± SD. ........................................................................................... 38 Figure 3.7 Compound class composition of wax mixtures from various B. distachyon organs. Waxes from cotyledons, spikes, leaf blades, leaf sheaths, stems and internal stems were analyzed. The coverage of each compound class is given as mean value in µg/cm2 ± SD. ............................................................................................................... 39 Figure 3.8 Chain length distributions within compound classes in waxes from various B. distachyon organs. Relative amounts of homologs are given as mean percentages of ix  the fraction within waxes from cotyledons, spikes, leaf blades, sheaths, stems, and internal stems (n = 6) ± SD. ......................................................................................... 41 Figure 3.9 Comparison of chain length distributions of individual wax components in green and etiolated leaf blades and sheaths. Percentages of individual homologs within the series of (A) fatty acids, (B) primary alcohols and (C) alkanes in green and etiolated leaf blades, and of (D) fatty acids, (E) primary alcohols and (F) alkanes in green and etiolated leaf sheaths are shown as mean values (n = 6) ± SD. ................................... 45 Figure 4.1 Possible positions of the methyl group in the new alkylresorcinol series. The methyl group may be located (A) on the resorcinol ring in the ortho position, (B) on the resorcinol ring in the para position, or else (C) in the (benzylic) position 1’ on the alkyl side chain. ........................................................................................................... 52 Figure 4.2 GC-MS single ion chromatograms (m/z 268) of TMSi-derivatized leaf blade waxes of S. cereale and B. distachyon. (A) Alkylresorcinols with C19 - C27 side chains were identified in wax from green leaf blades of S. cereale, and (B) with side chains of C17 - C25 for B. distachyon leaf blades. .................................................................... 56 Figure 4.3 Mass spectra of TMSi-derivatized alkylresorcinols in wax of S. cereale green leaf blades. (A) Alkylresorcinol with C19 side chain (AR 19:0; molecular ion m/z 520). (B) Alkylresorcinol with C21 side chain (AR 21:0; molecular ion m/z 548). (C) Alkylresorcinol with C23 side chain (AR 23:0; molecular ion m/z 576). (D) Alkylresorcinol with C25 side chain (AR25:0; molecular ion m/z 604). (E) Alkylresorcinol with C27 side chain (AR 25:0; molecular ion m/z 632). .................... 57 Figure 4.4 MS analysis of TMSi-derivatized C19:0 alkylresorcinols. (A) Mass spectrum of alkylresorcinol with C19 side chain (AR 19:0; molecular ion m/z 520) in wax from S. cereale green leaf blades, (B) synthetic standard of C19:0 alkylresorcinol, and (C) structure and fragmentation pattern of 5-n-nonadecylresorcinol................................. 58 Figure 4.5 Mass spectra of TMSi derivatives of methyl alkylresorcinols (MARs) in wax of green S. cereale leaf blades. (A) Methyl alkylresorcinol C19 (MAR 19:0; molecular ion m/z 534). (B) Methyl alkylresorcinol C21 (MAR 21:0; molecular ion m/z 562). (C) Methyl alkylresorcinol C23 (MAR 23:0; molecular ion m/z 590). (D) Methyl alkylresorcinol C25 (MAR 25:0; molecular ion m/z 618). ........................................... 61 Figure 4.6 Comparison of MS fragmentation patterns between methyl alkylresorcinols from S. cereale wax and organic synthesis. (A) Mass spectrum of C21 methyl alkylresorcinol (MAR 21:0; molecular ion m/z 562) in S. cereale green blade, and (B) mass spectrum of C21 methyl alkylresorcinol synthetic standard (CMAR 21:0; molecular ion m/z 562). ............................................................................................... 63 Figure 4.7 Comparison GC retention times between methyl alkylresorcinols from S. cereale wax and organic synthesis. (A) S. cereale green leaf blades sample showing four methyl alkylresorcinols (MAR) homologs C19, C21, C23 and C25, and C24 alkane internal standard. (B) Synthetic methyl alkylresorcinol (CMAR) with methylation in the 1’ position on the alkyl side chain, and C24 internal standard. .............................. 64 Figure 4.8 Chain length distributions of normal alkylresorcinols in the waxes of various S. cereale organs. (A) Green organs, (B) etiolated organs. The percentages of individual alkylresorcinol homologs are shown as mean values (n = 6) ± SD. ............................ 66 x  Figure 4.9 Alkylresorcinol chain length distributions in waxes on B. distachyon leaf blades and sheaths. The percentages of individual alkylresorcinol homologs are shown as mean values (n = 6) ± SD. ........................................................................................... 67 Figure 4.10 Chain length distributions of methyl alkylresorcinols in the total waxes on various S. cereale and B. distachyon organs. (A) Green S. cereale organs, (B) etiolated S. cereale organs, (C) leaf blades of B. distachyon. The percentages of individual methyl alkylresorcinol homologs are shown as mean values (n = 6) ± SD. ................ 68 Figure 4.11 Total alkylresorcinol amounts. (A) S. cereale organs in light and dark conditions, (B) B. distachyon organs. The coverages (µg/cm2) of alkylresorcinols are given as mean values (n=6) ± SD for the total wax extracted from entire organ.  Total alkylresorcinol amounts were determined by adding up quantities of all individual alkylresorcinol and methyl alkylresorcinol homologs detected by GC-FID. .............. 70 Figure 4.12 Semi-quantitative RT-PCR analyses of gene expression. (A) ScARS in S. cereale and (B) BdARS in B. distachyon. ScActin and 18S rRNA were used as constitutive controls in the analysis of ScARS and BdARS, respectively. Bl, blades; Hy, hypocotyl; Cy, cotyledon; Sh, sheath; exSh, external sheath; inSh, internal sheath; St, stem; iSt, internal stem; Sp, spike. ............................................................................................... 71 Figure 4.13 Subcellular localization of ScARS and BdARS. (A) C-terminal fusion of ScARS with GFP (35S:ScARS-sGFP) (B) C-terminal fusion of BdARS with GFP (35S:BdARS-sGFP); (C) and (D) the ER-specific marker 35S:HDEL-GFP were transiently expressed in N. benthamiana epidermal cells; (E) and (F) merged image. 73 Figure 4.14 Methyl alkylresorcinol potential biosynthetic pathways.  The biosynthesis of MAR using (A) one molecule of malonyl-CoA and two molecules of malonyl-CoA as extender in different sequences with straight fatty acyl-CoA as starter substrate; (B) three molecules of malonyl-CoA as extenders and one molecule of methyl-branched fatty acyl-CoA as starter substrate; (C) ARs as substrate for SAM-dependent methyl transferases forming ring-MARs. ................................................................................ 78 Figure 5.1 Chain length distributions of alkyl esters in the total waxes on various S. cereale organs. The percentages of individual alkyl ester homologs are given as mean values (n=6) ± SD. .................................................................................................................. 90 Figure 5.2 Chain length distributions of esterified alcohols in the waxes from various S. cereale organs. Waxes from green leaf blades, external and internal sheaths, etiolated leaf blades and sheaths were analyzed. Percentages of homologs are given as mean values (n=6) ± SD. ....................................................................................................... 92 Figure 5.3 Chain length distributions of free and esterified alcohols in the cuticular waxes on various S. cereale organs in light and dark conditions. Percentages of homologs are shown as means (n=6) ± SD. ....................................................................................... 97 Figure 5.4 Chain length distributions of free and esterified fatty acids in the cuticular wax of various S. cereale organs in light and dark conditions. Percentages of homologs are shown as means (n=6) ± SD. ....................................................................................... 99 Figure 6.1 Potential Cuticular wax biosynthetic pathways on grasses. The core represents C16 fatty acyl-CoA undergo successive elongations, extending the acyl chain by C2 units. xi  The VLCFAs wax precursors are then used for the production of aldehydes, primary alcohols and alkyl esters (left), and alkylresorcinols and alkanes (right). C26 primary alcohol is the central key compounds for alkyl esters formation. The biosynthesis of aldehydes is proposed as intermediate in the biosynthesis of primary alcohols. Cuticular alkylresorcinols are biosynthesized using VLCFAs as substrate. ............. 102                 xii  List of Abbreviations  9-BBN 9-borabicyclo[3.3.1]nonane ACP Acyl carrier protein ARs Alkylresorcinols ARSs Alkylresorcinol synthases  BSTFA  bis-N,O-(trimethylsilyl)trifluoroacetamide  CER  Eceriferum CHS  Chalcone synthase  CoA Coenzyme A dppf 1,1'-Bis(diphenylphosphino)ferrocene DGATs Diacylglycerol acyltransferases ECR  Enoyl-CoA reductase  ER Endoplasmic reticulum  EST  Expressed sequence tag  FAE Fatty acid elongase FID Flame ionization detector  GC Gas chromatography GFP  Green fluorescent protein  HCD β-hydroxylacyl-CoA dehydratase  KCR β-ketoacyl-CoA reductase  KCS β-ketoacyl-CoA synthase  LACS Long-chain acyl-CoA synthetase MAR Methyl alkylresorcinol MS  Mass spectrometry  NMR Nuclear magnetic resonance OLA Olivetolic acid PCR Polymerase chain reaction PKS Polyketide synthase POE Point of emergence xiii  SAM S-Adenosyl methionine SD Standard deviation STS  Stilbene synthase THF Tetrahydrofuran TMSi Trimethylsilyl  UV Ultraviolet VLCFA Very long chain fatty acid WSD Wax synthase                                 xiv  Acknowledgements  This research and thesis would not be possible without the support of many people. I would like to express my gratitude to my supervisor Dr. Reinhard Jetter, who has been the central key in my investigation, teaching me about science and nature. I will have eternal gratitude for his patience, support, knowledge, and his help with academic and personal issues.   Thanks to all the Jetter lab members, especially Luke, Daniela and Radu for their support, sharing time working together and having fun. To Mariya and Chen for their support, and June and Cassandra for bringing new air to our work place.   Deepest gratitude to my Chilean family, for giving me their support all the time, Rolando, Loreley, Liliana, Claudia, Cesar, Amanda e Ismael. Special thanks to my Canadian friends/Acadia Park community, whose have been important supports for me; Mike, Nora, Mark, Alejandro, Adriana, Angela, Derek, Raquel, Paolo and many others who are in my deepest heart.  Lastly, and the most important, thanks to my lovely wife Claudia and my sweet daughter Alfonsina, for boarding on this adventure together and be part of my life every day, every moment, and through every breath.      xv                    Para los amores de mi vida, Claudia y Alfonsina…  1  Chapter 1: Introduction 1.1 Plant cuticular waxes  1.1.1 The plant cuticle The plant cuticle is a lipophilic layer which covers the outer side of most areal plant organs. It is biosynthesized by epidermal cells and provides essential protection against abiotic and biotic stresses. One of the main functions of the cuticle is to prevent non-stomatal water loss, and this function is mainly exerted by the waxy components of the cuticle. For example, in tomato fruit aliphatic wax constituents form the main portion of the transpiration barrier (Vogg et al., 2004). The cuticle also plays further roles such as protecting against pathogens and ultraviolet (UV) radiation, and mediating interactions with insects (Eigenbrode & Jetter, 2002; Fürstner et al., 2005; Holmes & Keiller, 2002;). Those surfaces of plants covered with microcrystalline structures of surface waxes were demonstrated to have self-cleaning properties (Lotus-effect) protecting organs against dirt adhesion (Fürstner et al., 2005).  1.1.2 Structure and composition of the plant cuticle Plant cuticles consist of two major components defined by their solubilities in organic solvents (cutin and cuticular wax).  Within the cuticle, an outer layer called the cuticule proper and a more interior cuticular layer may be distinguished. The cuticle proper consists of cutin together with epicuticular wax deposited on top and intracuticular wax embedded within the cutin matrix. Thus, epicuticular wax forms the outermost layer of the cuticle, and it can occur in the form of wax films or wax crystals, depending on the plant species and 2  organs (Fig 1.1). The structure underneath of the cuticule proper is called the cuticular layer, which is composed of intracuticular wax, cutin and carbohydrates (Jetter et al., 2000).          Figure 1.1 Schematic cross-section of the plant epidermis and cuticle (Modified from Jetter et al., 2000).   The insoluble component of the cuticle is called cutin, a polymer consisting of ω- and mid-chain hydroxy and epoxy C16 and C18 fatty acids connected by ester bonds and also through glycerol bridges and contributing 40-80% of the cuticle mass (Heredia, 2003). In contrast, cuticular waxes are complex mixtures of very-long-chain fatty acids and their derivatives such as primary and secondary alcohols, aldehydes, alkanes, alkyl esters and ketones. Waxes also contain a wide range of cyclic compounds, including triterpenoids, flavonoids, phenylpropanoids and alkylresorcinols (Jetter et al., 2007). The abundance of specific wax components differs widely between species, organs and developmental stages of the same plant. For instance, Arabidopsis thaliana showed differences between the wax compositions on the top, middle, and base of the stems (Suh et al., 2005). More specifically, alkyl esters Intracuticular (wax + cutin) Epicuticular wax  (film and crystals)   Cuticule proper Cell wall Epidermal cells Cuticular layer  (wax + cutin + carbohydrates) 3  were reported to accumulate in certain organs or specific layers, for instance in Cosmos bipinnatus petals, where they were detected only in the adaxial cuticle (Buschhaus, 2010), or as seed storage reserves in jojoba (Simmondsia chinensis) (Benzioni & Forti, 1989).   1.1.3 Wax biosynthesis Plant cuticular waxes consist mainly of very-long-chain fatty acids (VLCFAs) and their derivatives. VLCFAs are biosynthesized first by elongation of C16 and C18 fatty acyl-CoAs, then by modification into various compound classes either on the alcohol-forming (reduction) or the alkane-forming (decarbonylation) pathways (Kunst et al., 2007). The enzymatic machinery responsible for wax biosynthesis has been studied mainly in Arabidopsis thaliana ( Jetter et al., 2006)  C16 and C18 fatty acyl units are biosynthesized de novo by the fatty acid synthase complex (FAS) in plastids of the epidermal cells. These long-chain fatty acids are exported to the endoplasmic reticulum (ER) via largely unknown mechanisms involving cleavage of the acyl carrier protein (ACP) by acyl-ACP thioesterase (Bonaventure & Salas, 2003; L Kunst et al., 2007) and esterification to coenzyme A (CoA) by a long-chain acyl-CoA synthetase (LACS). In the ER, the fatty acids are elongated in cycles of four reactions that extend the acyl-CoA by two carbons. Each cycle requires a fatty acid elongase (FAE) multi-enzyme complex formed by four different enzymes (Jetter et al., 2006; Kunst et al., 2007; Kunst & Samuels, 2009; Samuels et al., 2008; Wettstein-Knowles & Mikkelsen, 1984). The first reaction is catalyzed by a β-ketoacyl-CoA synthase (KCS) which condenses an acyl-CoA with malonyl-CoA to form β-ketoacyl-CoA. This is subsequently reduced to β-hydroxyacyl-CoA by β-4  ketoacyl-CoA reductase (KCR), then dehydrated to enoyl-CoA by β-hydroxyacyl-CoA dehydratase (HCD), and finally reduced to a saturated acyl-CoA by enoyl-CoA reductase (ECR) (Kunst & Samuels, 2003). The resulting VLC acyl-CoAs are further modified on one of two downstream biosynthetic pathways, either the acyl reduction pathway producing even-numbered primary alcohols and alkyl esters or the decarbonylation pathway producing odd-numbered compounds including alkanes, secondary alcohols and ketones (Kunst & Samuels, 2003). There is indirect evidence that in Arabidopsis thaliana aldehydes occur as intermediates of the alkane-forming pathway, prior to decarbonylation and therefore with even-numbered hydrocarbon chains. However, it has also been hypothesized that in other species aldehydes may rather be formed as intermediates along the alcohol-forming pathway (Jetter & Kunst, 2008)  1.2Alkylresorcinols  The large pools of VLC acyl-CoA intermediates present in epidermal cells suggest that they may serve for the biosynthesis of other compound classes as well, including the alkylresorcinols found in some plant species.  1.2.1 Occurrence and biological functions of alkylresorcinols 5-Alkylresorcinols (ARs) are a class polyketide-derived phenolic lipids (Fig 1.2), also called 1,3-dihydroxy-5-alkylbenzenes or 5-n-alkylresorcinols. Their amphiphilic properties are due to a combination of a hydrophobic alkyl chain and a resorcinol ring. Alkylresorcinols are 5  members of an extensive family of bioactive compounds, which have been identified in a large variety of plants and animals (Kozubek & Tyman, 1999).      Figure 1.2 Structure of 5-n-alkylresorcinol (n=1,2,3…).   ARs have been isolated from a variety of natural sources, which include eleven families of higher plants (Anacardiaceae, Ginkgoaceae, Proteaceae, Myrsinaceae, Primulaceae, Myristicaceae, Iridaceae, Compositae, Leguminosae and Gramineae) several non-vascular plants (algae, mosses), fungi and bacteria (Kozubek & Tyman, 1999). ARs occur as homologous series with alkyl chain lengths ranging from C5 to C29. For instance, the AR with a C15 side chain was initially found in Anacardium occidentale (Dawson, 1948), and later Ginkgo biloba was reported to contain C15 and C17 ARs (Zarnowska et al., 2000), while Mangifera indica ARs had C15,  C17 and C19 side chains (Knödler et al., 2007). Overall, it is important to note that the side chains of ARs have, in most cases, odd numbers of carbons. Further variation comes from branched or unsaturated side chains. In bacteria, very similar ARs had been detected in genera like Azotobacter, Mycobacterium and Pseudomonas (Funa et al., 2006), and in fungi such as Aspergillus, Corticium, Merulius, Neurospora, Phlebia, Phoma, Pulcherricium, Stemphylium, Streptomyces and Verticicladiella  (Jin & Zjawiony, 2006), among others.   6  ARs have a variety of biological properties including antifungal and antibacterial activities. For instance, ARs from Mangifera indica (Droby et al., 1986), Hordeum vulgare (García et al., 1997), Secale cereale (Reiss, 1989), Ginkgo biloba (Itokawa et al., 1989) and Lysimachia japonica (Arisawa et al., 1989) inhibit the growth of a range of pathogens and exhibit antitumor activities (Kubo et al., 1993; Arisawa et al., 1989). It was also reported that ARs have cytotoxic effects, for instance, against the A2780 ovarian cancer cell (Chaturvedula et al., 2002), most likely by causing changes in the biophysical properties of phospholipid bilayers including increased permeability (Stasiuk & Kozubek, 2010). All of these biological activities, together with the well-known roles of phenolic lipids as plant defense against a variety of pathogens and herbivores as well as various kinds of abiotic stresses (Kozubek & Tyman, 1999; Stasiuk & Kozubek, 2010), have led to the assumption that ARs play a pivotal role in plant protection.  The AR derivatives found in some plant species have important pharmaceutical properties. For instance, Sorghum spp. form 5-n-pentadecatrienyl resorcinol as an intermediate metabolite in the biosynthesis of sorgoleone, a benzoquinone found in exudates of root hairs (Baerson et al., 2008). In Cannabis sativa, the C5 AR olivetolic acid (OLA) serves as a first intermediate on the cannabinoid biosynthetic pathway (Taura et al., 2007; Taura et al., 2009).   Plant ARs have received attention for their food and nutrition values, due to their benefits on human health (Slavin, 2004). Numerous reports have shown that ARs occur at relatively high concentrations in cereal grains (Kulawinek & Kozubek, 2008; Ross et al., 2003), with highest values in wheat, rye and triticale (>500 µg/g), and relatively low levels on barley, millet, 7  maize, rice and sorghum (Chen et al., 2004; Ross et al., 2003). In cereals, ARs had been localized in the bran fraction, suggesting that ARs accumulate at or near the plant surface. Furthermore, ARs had also been detected in other non-edible organs such as seedlings of Oryza sativa, Secale cereale and Zea mays (Deszcz & Kozubek, 2000; Suzuki et al., 1996; Suzuki et al., 2003), and roots of Sorgum bicolor (Cook et al., 2010). However, the tissue localization of ARs were not assessed in these cases. On the other hand, high concentrations of ARs were found in leaves of Secale cereale (Ji & Jetter, 2008), and they were localized exclusively to the plant cuticular wax and hence the tissue surface.  ARs were also reported from Mangifera indica (mango) latex and peel (Droby et al., 1986), from Myristica fragrans seed covers, and from Anacardium occidentale shells (cashew nut) (Kozubek & Tyman, 1999), again indicating that ARs could be deposited in the cuticular waxes.   The surface localization of ARs suggests that they act as a first line of defense against pathogens and herbivores (Arisawa et al., 1989; García et al., 1997; Itokawa at al., 1989; Kubo et al., 1993; Reiss, 1989; Stasiuk & Kozubek, 2010). However, the real biological function of the surface ARs has not been determined. In order to test their role(s), it will be necessary to first understand the mechanisms underlying their formation and accumulation.  1.2.2   Biosynthesis of alkylresorcinols Similar to other phenolic compounds, plant ARs are biosynthesized by type III polyketide synthases (PKSs), which are the smallest and simplest PKS enzymes. PKSs produce a variety of aromatic products with single or multi-ring structures. Examples include stilbenes, pyrones, naphthalenes, phloroglucinols, chromones and curcuminoids (Rubin-Pitel et al., 8  2010). In all cases, PKSs catalyze reactions between acyl-CoA starters and malonyl-CoA extenders, leading to a series of decarboxylative condensation cycles. Chalcone synthase (CHS) and stilbene synthase (STS), which are involved in naringenin chalcone and stilbenoid biosynthesis, respectively, were the first type III PKSs discovered. CHSs and STSs both use aromatic acyl CoA substrates and carry out three extensions, however they differ in their cyclization mechanisms (Fig 1.3) (Austin & Noel, 2003; Baerson et al., 2010; Cook et al., 2010). CHS is involved in the biosynthesis of flavonoids that are important for flower pigmentation (Winkel-Shirley, 2001), plant defense  (Cushnie & Lamb, 2005) and ultraviolet (UV) photoprotection (Winkel-Shirley, 2002). Other type III PKS enzymes differ in their substrate preferences, the number of extension reactions, and/or the cyclization mechanism.           Figure 1.3 Chalcone synthase and stilbene synthase mechanism for forming naringenin chalcone and resveratrol using Claisen and aldol condensations, respectively.   Based on this knowledge, it was hypothesized early on that ARs are biosynthesized by specialized PKS enzymes, AR synthases (ARSs), analogous to STSs but using aliphatic HOSCoAOHOOS-EnzO O O HOOHOHHOOHOOHOH3 x malonyl-CoAp-coumaroyl-CoA tetraketide intermediateresveratrolnarigenin chalconetype III PKSSTSCHS9  instead of aromatic acyl-CoA starters (Fig 1.4). Three sequential decarboxylative condensation reactions of the fatty acyl-CoA starter with three molecules of malonyl-CoA should lead to a tetraketide that undergoes aldol condensation to yield the AR product (Austin & Noel, 2003; Funa et al., 2006; Funabashi et al., 2008). The biosynthesis of ARs had been investigated experimentally in etiolated rice seedlings (Suzuki et al., 2003), showing that fatty acid units indeed act as direct precursors and form the side-chain moiety of 5-n-alkylresorcinols, and that the ARS enzymes exhibit substantial substrate specificity.       Figure 1.4 Possible mechanism for the formation of alkylresorcinols through elongation and aldol condensation catalyzed by alkylresorcinol synthase (ARS).   The mechanisms described above were initially confirmed in vivo and using (partially) purified ARSs. Later, corresponding enzymes from diverse organisms were isolated and further characterized. One of the first ARSs identified was ArsB from the bacterium Azobacter vinelandii (Funa et al., 2006), which is responsible for n-heneicosylresorcinol (AR21:0) formation. Also, an SrsA from Streptomyces griseus (Funabashi et al., 2008) was identified, which uses branched C15 and straight C16 acyl-CoAs as starter substrates and both malonyl-CoA and methylmalonyl-CoA as extender units, resulting in methyl-branched ARs. Besides the ARSs found in bacteria, ARSs from plants have been characterized. Most notably, two ARSs from S. bicolor, with in vitro activities on fatty acyl-CoA starter substrates, are forming intermediate alkylresorcinols for sorgoleone biosynthesis (Cook et al., 3 x malonyl-CoAtype III PKSH3C SCoAOn H3COn S-EnzO O OH3COHOHnARSCO2fatty acyl-CoA tetraketide intermediates alkylresorcinols10  2010). In Cannabis sativa, olivetol synthase (OLS) is involved in the biosynthesis of the short-chain alkylresorcinol (Taura et al., 2009). All plant ARSs characterized to date thus have substrate preferences for acyl-CoA starters with short to long chains, but not VLC acyls. However, the ARs found in cereals have very long side chains, suggesting that ARSs involved in their biosynthesis must accept VLCFA starter substrates (Ji & Jetter, 2008). In order to test this hypothesis, ARSs responsible for the biosynthesis of cuticular alkylresorcinols must be investigated.  1.2.3 Secale cereale and Brachypodium distachyon as model system Previous members of our group have chosen both S. cereale and B. distachyon as model systems for investigating the biosynthesis of cuticular ARs. Both species are part of the Pooideae subfamily within the Gramineae, an important clade providing the bulk of human food. Secale cereale had been reported to contain the highest AR amounts of all cereal grains (Ross et al., 2004). Additionally, B. distachyon was further chosen as the first member of the Pooideae to have its genome sequenced, and there are many genetic tools currently being developed for this species. Also, B. distachyon presents diverse advantageous biological attributes as a model organism, such as self-pollination, fast growth, a small genome (~275 Mb harboring 26,000 genes excluding isoforms), and a close phylogenetic relation to a large and diverse group of temperate cereal crops (Alves et al., 2009)  The composition of very-long-chain alkylresorcinols in leaf waxes of Secale cereale had been investigated, indicating that AR homologs occur at relatively high levels when the second leaves of plants are 20 cm long (15 - 17 days old). It was also reported that cuticular 11  wax on B. distachyon leaves contains substantial amounts of ARs, albeit with slightly shorter chain lengths than rye leaves (Yao, 2011). In addition, methylated alkylresorcinols had been found on B. distachyon previously in our lab (Yao, 2011). Similar compounds were also described once in wheat with chain lengths ranging from C19 to C27 (Adamski et al., 2013). In both reports, the methyl alkylresorcinol structures were inferred from the MS fragment m/z 282, however the methylation position could not be determined yet.  1.2.4 Isolation and partial characterization of BdARS and ScARS Two genes potentially involved in the biosynthesis of cuticular alkylresorcinol had been isolated from Secale cereale (ScARS) and B. distachyon (BdARS) (Fig 1.5) and partially characterized (Yao, 2011). In B. distachyon, gene cloning was performed based on the mining of B. distachyon expressed sequence tag (EST) libraries. In Secale cereale, PCR-based gene cloning was carried out using primer designs exploiting homology between the functionally characterized ARS sequences from Sorghum bicolor and Oryza sativa (Yao, 2011). Both ScARS and BdARS share relatively high amino acid similarity with PKSs such as Gh2PS from Gerbera hybrida, MsCHS from Medicago sativa, OsARSs from Oryza sativa, ScCHSs from Secale cereale, and SbARSs from Sorghum bicolor (Yao, 2011).      12        Figure 1.5 Gene structures of previously isolated BdARS from B. distachyon and ScARS from S. cereale. Exons are shown in shaded boxes and introns are shown in lines (Adapted from Yao, 2010).  The presence of residues putatively associated with the catalysis and CoA binding and of residues Cys164, His303 and Asn336, representing the catalytic triad conserved in all plant type III PKSs, was of special interest. In addition, in vivo tests using heterologous expression of ScARS and BdARS in two different yeast strains demonstrated that both proteins had ARS activity (Yao, 2011). However, the AR homologs formed had shorter chains than those in rye and B. distachyon waxes. It was therefore not clear whether the ARSs are indeed involved in formation of the cuticular ARs. In order to further test involvement of the genes/proteins in the formation of cuticular ARs, they had to be characterized further.  As one approach to test the involvement of ScARS and BdARS in cuticular AR biosynthesis, the expression patterns of both genes had to be compared with product accumulation profiles. Accordingly, a partial gene expression study of the ARS genes in various plant organs had previously been performed in our lab (Yao, 2011). Different organs of Secale cereale (green leaf blade, etiolated leaf blade, cotyledon, leaf sheath and root) and B. distachyon (green leaf, 13  stem, spike and root) were examined to test for ARS gene expression. However, this study was not completed due to long growth times of some plant organs, and the expression-product correlation therefore remained inconclusive. A more comprehensive gene expression analysis is necessary to determine whether both BdARS and ScARS are indeed expressed at the predicted time and place for ARs formation.  1.2.5   Effects of etiolation on cuticular alkylresorcinols It is well established that light exposure affects de novo fatty acid synthesis in chloroplasts (Sasaki et al., 1997). Consequently, it was also hypothesized that light levels during organ growth may have effects on wax formation. In Brassica oleracea, it was accordingly found that the biosynthesis of hydrocarbons and secondary alcohols increased at the expense of aldehydes and esters in bright light conditions (Macey, 1970).  Also, the amounts of wax per leaf surface area were affected by the light and temperature in expanding leaves of Hordeum vulgare (Giese, 1975). All taken together, it was concluded that light stress affected fatty acid precursor amounts.   If cuticular ARs are indeed formed from the same acyl-CoA intermediates as VLCFA derivatives, then etiolation should have similar effects on ARs as on other wax constituents. In accordance with this hypothesis, a series of homologous alkylresorcinols 13:0, 15:1, 15:0, 17:1, 17:0 were found in etiolated rice seedlings (Suzuki et al., 1996), indicating that ARs side chain length profiles may indeed be changed when plants are grown in the dark. Furthermore, the total amount of ARs in etiolated rice seedlings were higher than in green 14  rice seedling, and AR amounts decreased rapidly when the etiolated seedlings were exposed to light (Suzuki et al.,1996). Therefore, etiolation provides an important additional tool to bring about changes in AR levels and to correlate them with gene activity changes, and also to determine whether the variation of VLCFA CoA pools is correlated with AR side chain length profiles.   1.3 Research questions and objectives The current study was based on the hypothesis that ScARS and BdARS, previously isolated from Secale cereale and B. distachyon, respectively, are enzymes involved in the biosynthesis of cuticular very-long-chain alkylresorcinols, by preferentially accepting VLC acyl-CoAs precursors as substrates that accumulate in ER membranes of epidermal cells during wax production.   In order to test this hypothesis, various experiments were performed using analytical techniques to identify and quantify wax components in a suite of plant organs. One main objective was to analyze the main components in the cuticular waxes of Secale cereale and B. distachyon organs (chapter 3). To this end, plants grown in the light or in the dark were compared, to test the effect of etiolation on compound class distributions and chain length profiles. A second major objective was to analyze alkylresorcinol homologs in the cuticular waxes of Secale cereale and B. distachyon, and to compare their amounts with ScARS and BdARS gene expression levels in various organs (chapter 4). In addition, also the subcellular localization of corresponding proteins was studied to test whether indeed ARS proteins have access to VLC acyl-CoA intermediates of wax biosynthesis. Finally, a third objective was to 15  identify and quantify alkyl ester homologs and isomers within the wax mixture on various organs of Secale cereale, to provide indirect evidence for the chain length distribution of the acyl-CoA pools accumulating during wax biosynthesis.                 16  Chapter 2: Materials and methods 2.1 Plant material and growth conditions 2.1.1 Plant growth conditions Seeds of Secale cereale L. cv. Esprit were purchased from Capers, Vancouver, Canada, and grains of Brachypodium distachyon (L.) P. Beauv. were ordered from National Germplasm Resources Laboratory, Beltsville, Maryland, United States of America. Seed of Nicotiana benthamiana were provided by Dr. Mathias Schuetz (Samuels lab, Department of Botany, University of British Columbia). Seeds of S. cereale and B. distachyon were soaked in water at room temperature overnight. Then, they were planted in plastic pots (diameter 15 cm) containing moistened Sunshine Mix #4 (Sun Gro Horticulture Canada). Seeds of N. benthamiana were sown directly in pots containing moistened Sunshine Mix #4. All of them were stratified in the dark for 2 days at 4°C and then moved to a growth chamber at The University of British Columbia, where they were kept for 20 hours at 22°C in the light and for 4 hours at 18°C in the dark, at 70% of relative humidity. Etiolated plants were grown in dark conditions for 16 hours at 22°C and 8 hours at 18°C in 70% humidity.  2.1.2 Plant materials After three weeks of growth, plants were harvested and dissected. Secale cereale etiolated organs selected were leaf blades, sheaths, stems, hypocotyls and cotyledons. In the case of S. cereale plants grown in light conditions, leaf blades and sheaths were harvested. Two different types of sheaths were distinguished as external sheaths exposed directly at the plant surface, and internal sheaths enclosed within sheaths of older leaves. In B. distachyon, only 17  green materials were collected because specimens could not be grown reproducibly in the dark. Organs selected were leaf blades, sheaths, cotyledons, spikes and stems. In this particular case, stems were divided in two portions; the part of the stems exposed directly at the surface of the plant, and internal stem parts embedded inside of a leaf sheath.   The wax extraction was carried on immediately after the harvest and the dissection of plant organs. Six independent parallels were analyzed in all green organs.  In the case of etiolated material, 13 collected organs were pooled together into one sample, and then six pooled samples were analyzed independently. For semi-quantitative RT-PCR analysis, all organs selected in both conditions were harvested separately and stored at -80 C for posterior analysis.   For subcellular localization, 4-week-old leaves of Nicotiana benthamiana were used for agrobacterium-mediated infiltration experiment. Plants were returned into the growth chamber under the same conditions described before for 3-4 days. Then, transformed leaves were examined using confocal microscopy.  2.2 Wax analysis 2.2.1 Wax extraction and derivatization Selected organs were harvested and photographed with a digital camera for area analysis using the program Image J (http://imagej.nih.gov/ij/). In leaf blades, sheaths, hypocotyls and cotyledons, the area calculated were multiplied by two, representing the wax extracted on both abaxial and adaxial face. In the case of stems, the area calculated were multiplied by π. 18  Total wax was extracted by immersing the entire organs separately twice for 30 seconds in chloroform containing defined amounts of n-tetracosane and 5-n-tridecylresorcinol as internal standards. The solutions were concentrated under a gentle stream of N2 gas while heating to 50°C and transferred to sample vials. Subsequently, waxes were derivatized with bis-N,O-(trimethylsilyl) trifluoroacetamide (BSTFA; Sigma-Aldrich) in pyridine (1:1, v/v) at 70°C for 40 min to transform all hydroxyl- containing compounds into the corresponding trimethylsilyl (TMSi) derivatives. The solvent was evaporated under a stream of nitrogen gas, and then chloroform was added to the sample prior to quantitative analyses by gas chromatography-flame ionization detection (GC-FID) and compound identification by gas chromatography-mass spectrometry (GC-MS).  2.2.2 Chemical analysis by GC-FID and GC-MS Qualitative analyses of the wax compositions were performed using capillary GC (5890N, Agilent, Avondale, PA; column 30 m HP-1, 0.32 mm i.d., df=0.1 µm, Agilent) with temperature-programmed on-column injection at 50°C, separation using a temperature program for 2 min at 50°C, raised by 40°C per minute to 200°C, held for 2 min at 200°C, raised by 3°C min-1 to 320°C and held for 30 min at 320°C and He carrier gas inlet pressure programmed for a constant flow of 1.4 ml/min. Wax components were identified with a mass spectrometric detector (5973N, Agilent) by comparison of characteristic fragmentation patterns with those of authentic standards and literature data. Quantitative analyses were carried out by GC using a temperature program as described above, but with a flame ionization detector (FID) and H2 carrier gas inlet pressure programmed for a constant flow of 2.0 ml/min. Each GC-FID peak was identified using GC-MS chromatograms, then integrated 19  and compared with that of an internal standard n-tetracosane added at the wax extraction procedure. Finally the values were divided by the surface area of the sample to obtain measurements in µg/cm2. All quantitative data are given as means of parallel experiments and standard errors.  2.2.3 Synthesis of C19 methyl alkylresorcinol (CMAR19:0) The synthesis of C19 CMAR (methyl alkylresorcinol with methylation in position 1’ on the alkyl side chain) was performed using the same method described in Ji, 2010 (Fig 2.1). A mixture of 3,5-dimethoxyphenol (1.5 g, 9.8 mmol, Sigma–Aldrich) and 2,6-lutidine (1.6 ml, 13.7 mmol, Sigma–Aldrich) in CH2Cl2 (48 ml) at 10°C was added slowly to a solution of triflic anhydride (2.1 g, 7.4 mmol, Sigma–Aldrich) in CH2Cl2 (9 ml). The mixture wax was cooled to 0°C and stirred for 2 hours. The organic layer was separated and dried with Na2SO4, and the solvent evaporated under vacuum. The product was purified by column chromatography using packed silica gel and hexane:ethyl acetate (10:1) as mobile phase. Fractions were dried overnight under a stream of nitrogen gas, and 3,5-dimethoxyphenol triflate was obtained with 45% of yield. Structure was confirmed by NMR. Separately, 1-eicosene (401 mg, 0.7 ml, 2.6 mmol, Signa-Aldrich) and 9-borabicyclo[3.3.1]nonane (9-BBN) (5.2 ml, 2.6 mmol, Sigma–Aldrich) in tetrahydrofuran (THF) (60 ml) were stirred for 2 hours under N2 at room temperature. After that, NaOMe (0.17 g, 3 mmol), PdCl2 (dppf) (56 mg, 0.07 mmol, Sigma–Aldrich) and 5-dimethoxyphenol triflate synthesized before (0.65 g, 2.3 mmol) were added, the mixture heated and allowed to reflux for 1 hour, the solvent evaporated and CH2Cl2 (10 ml) added. The product was purified through a short column of 20  silica removing the crude product by flash CC with hexane/EtOAc (10:1), to form 1,3-dimethoxy-5-nonadecylbenzene.                       Figure 2.1 Synthesis of C21 methyl alkylresorcinol with methylation on the benzylic carbon. (*) Represent the minor products synthesized in the reaction.   B HOCH3H3CO OHF3C S OOOS CF3OON CH3H3COCH3H3CO O S CF3OOCH2Cl2, 0 C, 2hn=8Bn=8OCH3H3CO O S CF3OOOCH3H3COB IOHHOn=8n=8++++THF, room temp, 2hTHF, PdCl2 (dppf), reflux, 1hHexane, room temp, 4hBn=8OCH3H3COn=8Bn=8Bn=8OCH3H3COn=8OCH3H3COn=8OHHOn=8+++++** ***21  Finally, 9-iodo-9-BBN (0.41 ml, 2.52 mmol, Sigma–Aldrich) and the 1,3-dimethoxy-5-nonadecylbenzene synthesized before (0.5 g, 1.2 mmol) were mixed with hexane (25 ml) and stirred for 3 hours at room temperature. Following solvent evaporation under vacuum, 15 ml of Et2O was added to dissolve the residue. The 9-BBN ethanolamine product was precipitated with ethanolamine (0.14 ml, 2.2 mmol, Sigma–Aldrich) in THF (1 ml), and the mixture was stirred for 3 hours. The precipitate was removed with filter paper and the filtrate was brought to dryness. Finally, flash CC was utilized to purify the product using hexane/EtOAc (2:1) as eluent. The products were confirmed mediate NMR procedure.   2.3 Genetic analysis 2.3.1 RNA isolation and reverse transcription  Three green (leaf blades, external leaf sheaths and internal leaf sheaths) and five etiolated (leaf blades, leaf sheaths, stem, hypocotyl and cotyledon) S. cereale organs, and six green B. distachyon organs (cotyledons, spikes, leaf blades, leaf sheaths, stems and internal stems) were harvested (2.1.2), and RNA was extracted from them using RNeasy Plant Mini Kit (Qiagen). During RNA extraction, samples were treated by RNase-Free DNase (Qiagen) to avoid DNase digestion. The amounts of extracted RNA were measured using a NanoDrop 8000 Spectrophotometer (Thermo Scientific). Two micrograms of total RNA were used to perform the reverse transcription with oligo (dT) primer for first-strand cDNA synthesis by SuperScript II Reverse Transcriptase (Invitrogen) at 42°C for 60 min. The resulting cDNA was subsequently used as template in PCR reactions.  22  2.3.2 Semi quantitative RT-PCR ScARS gene-specific forward primer 5’-AAGCATAGGAACCACCAACGGCAA-3’ and reverse primer 5’-AAGACGAGGTGGGTGATCTCGCT-3’ were used to amplify and detect gene expression in etiolated and green S. cereale organs.  ScActin was amplified as a positive control using ScActin gene-specific forward primer 5’-ATGCTAGTGGACGCACAACAGG TA-3’ and reverse primer 5’-ATCTTCATGCTGCTTGGTGCAAGG-3’. In gene expression analysis of ScARS, RT-PCR was carried out under the conditions: 98°C for 30 s, 26 cycles of 98°C for 15 s, 63°C for 30 s, and 72°C for 30 s, and 72°C for 5 min with Phusion High-Fidelity DNA Polymerase.  Specific forward primer 5’-CGACCAGTTCTTCCGCGTGACC-3’ and reverse primer R 5’-GATGGCTGGTCTCGGTCGAGGA-3’ were used to amplify a fragment of BdARS using cDNA templates from various B. distachyon organs. For positive control, specific forward primer 5'-CCGTCCTAGTCTCAACCATAAAC-3' and reverse primer 5'-CCTTTAAGTTTCAGCCTTGCG-3’ were used to amplify 18S rRNA. RT-PCR was carried out under the conditions: 98°C for 30 s, 24 cycles of 98°C for 15 s, 60°C for 30 s, and 72°C for 30 s, and 72°C for 5 min with Phusion High-Fidelity DNA Polymerase.   2.3.3 Agrobacterium-mediated infiltration To investigate the subcellular localization of BdARS and ScARS, the GFP fusion constructs prepared by former Master student Rounan Yao were used to transform Agrobacterium tumefaciens strain GV3101 according to a previous protocol (Sparkes et al., 2006). A. tumefaciens containing plasmids with the construct 35S:sGFP-BdARS/ScARS or 23  35S:BdARS/ScARS-sGFP were grown at 28°C in LB medium with kanamycin (50 µg/ml), hygromycin (50 µg/ml), rifampicin (100 µg/ml) and gentamicin (50 µg/ml) overnight. Pellets containing cells were collected by centrifugation at 5000 rpm for 15 min at room temperature and washed with dH2O, and resuspended in 4 ml of dH2O. The abaxial air spaces of three-week-old N. benthamiana leaves were infiltrated with the Agrobacterium tumefaciens suspension using a sterile syringe. The ER-specific control marker line p35S:HDEL-RFP was co-infiltrated with each construct. Infiltrated plants were placed back into the growth chamber under the same conditions informed before (2.1.1). After three days, transgenic N. benthamiana leaves were harvested and analyzed using an Olympus multi-photon confocal microscope (BioImaging Facility, Department of Botany, University of British Columbia) with an excitation of 488 nm for GFP and 543 nm for RFP. Then, images obtained were processed using Volocity software.            24  Chapter 3: Total wax profiles in various organs of Secale cereale and Brachypodium distachyon 3.1 Introduction It is well known that plant waxes are complex mixtures of very-long-chain aliphatics such as alkanes, primary and secondary alcohols, aldehydes, ketones, fatty acids and esters. Due to their roles in protecting the plant tissue against adverse, changing environments, it has long been surmised that plant waxes may be formed, at least in part, in response to stress. It has been shown that both the composition and total amounts of wax mixtures vary depending on growth conditions for some species. Most notably, drought stress was found to affect wax composition in Arabidopsis thaliana (Kosma et al., 2009) where the increase in alkane chain lengths was observed, or on roses (Jenks et al., 2001), on Abutilon theophrasti (Levene & Owen, 1995), and on varieties of  Sorghum (Premachandra et al., 1992) elevated wax loads had been reported under limited water conditions. Changes in temperature also has been reported to affect wax accumulation, in Brassica oleracea (Baker, 1974)  and Brassica napus (Whitecross & Armstrong, 1972) the decrease in temperature induces the largest deposits of wax and changes in the ultrastructure of the wax surface.   It had also been reported that stress exerted by low light levels during growth affected plant wax amounts and composition. In particular,  etiolated  leaves of Brassica napus (Whitecross & Armstrong, 1972) the effect of reduction in light intensity showed lower levels of wax deposition, and Brassica oleracea (Macey, 1970; Baker, 1974) were found the light altered the balance of synthesis of the various wax components. Many other investigations on wax compositions of diverse species did not vary growth conditions, and it remains to be tested 25  whether dynamic adaptation of wax composition is restricted to a few species or a more general phenomenon. Therefore, the purpose of the present study was to test the effects of etiolation on wax composition through comprehensive analyses of waxes from various organs of selected model species.   Monocots present a particularly interesting case for studying light effects on wax deposition, based on ample but indirect evidence from organ comparisons. Previous investigations showed that leaf blades had drastically higher wax coverages than sheaths, suggesting that wax deposition occurred mainly at or near the point of emergence (POE) between sheath and blade. It was therefore concluded that during leaf epidermis development of grasses exposure to light at the POE was an important trigger for wax formation. It was hypothesized that etiolation of entire grass plants should also result in changes of wax amounts and possibly composition. However, this has to date been tested only on one monocot species, Hordeum vulgare (Giese, 1975). In this case, wax amount and chain length dominated had variation in plants grown in light and the dark, but only limited chemical information was provided at the time. Therefore, the goal of the present work was to carry out comprehensive analyses of waxes from various organs, grown in the light or in the dark, of selected grass species.  Secale cereale and Brachypodium distachyon have recently emerged as model systems in the Poaceae due to several reasons: i) S. cereale provides the bulk of human nutrition and; ii) this specie has been reported to contain the highest alkylresorcinol amounts of all cereal grains (Ross et al., 2004); iii) B. distachyon is the first member of the Pooideae subfamily to have its genome sequenced, and; iv) presents diverse advantageous biological attributes, such as 26  self-pollination, fast growth, a small genome (~275 Mb harboring 26,000 genes excluding isoforms) and a close phylogenetic relation to a large and diverse group of temperate cereal crops (Alves et al., 2009); and finally v) according to the phylogenetic relationship, S. cereale and Brachypodium distachyon are closely related, being both temperate cereal species.   Some chemical, biological and biochemical investigations into cuticular wax in S. cereale and Brachypodium distachyon had previously been performed in our lab. The total wax mixtures from both sides of the leaves of S. cereale contained primary alcohols (71%), alkyl esters (11%), aldehydes (5%), and small amounts (<3%) of alkanes, steroids, secondary alcohols, fatty acids, and novel homologous series of alkylresorcinols (3%) (Ji & Jetter, 2008). Brachypodium distachyon leaf wax was found to consist of primary alcohols (71%), alkyl esters (11%), aldehydes (2%), and alkanes (2%), and alkylresorcinols (5%) (Yao, 2011). In further, more detailed studies, the distribution of waxes along the length of S. cereale leaf blade and their accumulation over time were assessed. The major compound classes were all found to accumulate primarily in the same zone of the leaf at/beyond the POE (Ji, 2010). In addition, the same study showed that leaf one had very small amounts of alkylresorcinols (ARs) while leaves two, three and four all had similar, higher AR coverages. Based on all this information, leaf two was chosen for the present investigations into the effect of etiolation on sheath and blade waxes.  Overall, the goal of this chapter was to analyze the chemical composition of the cuticular wax of five etiolated and three green (non-etiolated) S. cereale organs, and six green 27  Brachypodium distachyon plant organs. Using gas chromatography-flame ionization detector (GC-FID) and gas chromatography-mass spectrometry (GC-MS), total waxes were quantified and identified by compound class in the current chapter, addressing the following question: How is the wax profile distribution in different plant organs? Are there wax composition differences in etiolated and non-etiolated plant organs? The results will not only provide insights on light effects on wax deposition, but also provide the context for more detailed analyses of ARs and alkyl esters, and a deeper understanding of the biosynthesis of these compound classes (chapter 4 and 5).                28  3.2 Results  The overall goal of this chapter was to identify and quantify the components of cuticular waxes on various green and etiolated S. cereale and Brachypodium distachyon organs. To this end, cuticular waxes from green S. cereale leaf blades, external and internal sheaths, from etiolated S. cereale leaf blades, sheaths, stems, hypocotyls and cotyledons (3.2.1), and from green Brachypodium distachyon leaf blades, stems exposed to the light and enclosed within sheaths, cotyledons, spikes and sheaths (3.2.2) were extracted and analyzed by GC-FID and GC-MS.    3.2.1 Cuticular wax profiles of various S. cereale organs The total wax mixtures were extracted from selected entire organs, without distinguishing wax locations in the epicuticular and intracuticular layers or the abaxial and adaxial leaf sides. In order to assess the overall effect of light stress, growth was monitored for etiolated and for normally grown plants (3.2.1.1), total waxes were quantified from various green and etiolated organs of S. cereale (3.2.1.2), together with compound class compositions (3.2.1.3) and their respective chain length distributions (3.2.1.4).   3.2.1.1 Growth of S. cereale leaves in the light and in the dark To assess how light stress impacts leaf growth, the size of leaf two was monitored by measuring the length of the exposed leaf blades and sheaths as a function of time (Fig 3.1). Leaf blades and sheaths were distinguished as the portions of the leaves above and below the point of emergence (POE) the point where adaxial epidermis cells become exposed.   29  Green leaf blades grew from a length of 2.6 cm on day 6 to 19.5 cm on day 16, with a growth rate of ca 1.7 cm/day. After day 16, green leaf blades further grew to a length of 22.5 cm, with a growth rate of ca 0.75 cm/day, resulting in an overall average growth of 1.4 cm/day over the entire monitoring period. In the dark, leaf blades grew from 3.8 cm on day 6 to 9.4 cm on day 12, with a growth rate of ca 1 cm/day, and then to 10.9 cm maximum length at day 20 with drastically decreased growth of ca 0.18 cm/day. Overall, etiolated leaf blades grew on average 0.5 cm/day during the 20 days of measurement. Green sheaths grew from 0.9 cm on day 6 to 5.5 cm on day 16, with a growth rate of ca 0.46 cm/day. At day 20, green sheaths reached a length of 6.5 cm, showing a growth rate of 0.25 cm/day in the last four days, resulting in an overall average of 0.4 cm/day. Etiolated sheaths grew very slowly from 1.2 cm on day 6 to 3.5 cm on day 14, with a growth rate ca 0.29 cm/day. After day 14, etiolated sheaths had a growth rate of only 0.07 cm/day, for an overall average growth of 0.2 cm/day.           30              Figure 3.1 Growth of green and etiolated S. cereale leaf blades and sheaths. Blades and sheaths were distinguished as the parts of the leaf above and below the POE (point of emergence). Growth was monitored by measuring blade and sheath length in situ until day 20 after germination. Data are given as mean values (n=4) ± SD.   3.2.1.2 Total wax amounts on S. cereale organs To analyze waxes, they were extracted from various parts of S. cereale plants by submerging them in chloroform, transformed into trimethylsilyl (TMSi) derivatives, and then identified using GC-MS and quantified by GC-FID. Total wax coverages were determined for three green (leaf blades, external sheaths and internal sheaths) and five etiolated (leaf blades, sheaths, stems, hypocotyls and cotyledons) S. cereale organs.  Green leaf blades (16.2 ± 2.9 µg/cm2) had the highest wax coverage of all S. cereale organs selected (Fig 3.2). In comparison, green external sheaths (6.9 ± 1.4 µg/cm2) had ca 43% of Time [days after germination]6 8 10 12 14 16 18 20Length [cm]0510152025Green bladesEtiolated bladesGreen sheathsEtiolated sheaths31  the wax load of green leaf blades, and green internal sheaths (5.1 ± 0.3 µg/cm2) had ca 32% of the load. Etiolated leaf blades (12.1 ± 2.7 µg/cm2) had ca 75% of the wax load of green leaf blades. Etiolated sheaths (2.9 ± 0.4 µg/cm2) had ca 25% of the wax amount of etiolated leaf blades. All other etiolated organs had intermediate wax loads: stems 4.8 ± 0.5 µg/cm2, hypocotyls 4.2 ± 0.9 µg/cm2, and cotyledons 3.2 ± 0.7 µg/cm2.               Figure 3.2 Total wax loads on various S. cereale organs grown in the light or in the dark. Wax coverages are given as mean values (n = 6) ± SD.  3.2.1.3 Compound class distribution in waxes on etiolated and green S. cereale organs Waxes from various S. cereale organs comprised typical compound classes, including fatty acids, primary alcohols, alkanes, aldehydes and alkyl esters together with alkylresorcinols Green leaf bladesEtiolated leaf bladesGreen ext. sheathsGreen int. sheathsEtiolated sheathsEtiolated stemsEtioalted hypocotylsEtiolated cotyledonsWax coverage [µg/cm2 ]0510152032  (Fig 3.3). The relative amounts of these compound classes differed between organs and also depending on growth in the light or in the dark. Green leaf blades had primary alcohols as the major wax constituents, representing ca 65% of the total wax (10.6 ± 2.1 µg/cm2), together with ca 11% aldehydes (1.8 ± 0.5 µg/cm2), ca 10% alkyl esters (1.6 ± 0.4 µg/cm2), ca 3% alkanes (0.5 ± 0.1 µg/cm2), ca 2% alkylresorcinols (0.3 ± 0.05 µg/cm2), ca 0.8% fatty acids (0.1 ± 0.02 µg/cm2), and ca 7.2% other minor compounds plus non-identified compounds (1.2 ± 0.4 µg/cm2). The other two green organs (external and internal sheaths) had similar patterns, with primary alcohols as predominant compound class (2.5 ± 0.7 µg/cm2 and 43% 2.2 ± 0.1µg/cm2, respectively). However, both organs had varying amounts of other compound classes, in external sheaths 1.2 ± 0.1 µg/cm2 aldehydes, 0.9 ± 0.1 µg/cm2 alkyl esters, 0.5 ± 0.03 µg/cm2 alkanes, 0.4 ± 0.04 µg/cm2 alkylresorcinols and 0.1 ± 0.05 µg/cm2 fatty acids. Wax from internal sheaths contained the same compound classes in slightly different ratios.  Waxes from etiolated leaf blades and stems had similar compound class compositions, in which primary alcohols were the predominant compound class with 61% (7.4 ± 2.1 µg/cm2), and 36% (1.8 ± 0.2 µg/cm2), respectively. Alkyl esters were the second most abundant group of compounds with 6% (0.7 ± 0.2 µg/cm2) and 10% (0.5 ± 0.06 µg/cm2), respectively. Alkylresorcinols represented 2% and 8% of the waxes of both organs, respectively, followed by fatty acids (1% and 7%), and alkanes (1% and 2%).     33             Figure 3.3 Compound class compositions of wax mixtures from various S. cereale organs. Waxes from green leaf blades, external sheaths and internal sheaths, and from etiolated leaf blades, stems, hypocotyls, cotyledons and sheaths were analyzed. The coverage of each compound class is given as mean value in µg/cm2 ± SD.     Etiolated hypocotyls and leaf sheaths also had primary alcohols as the most abundant compound class with 17% (0.8 ± 0.1 µg/cm2) and 32% (1.0 ± 0.1 µg/cm2), respectively. Hypocotyl wax contained 17% alkyl esters (0.7 ± 0.1 µg/cm2), 16% fatty acids (0.6 ± 0.1 µg/cm2), 3% alkanes (0.1 ± 0.03 µg/cm2) and 2% alkylresorcinols (0.1 ± 0.01 µg/cm2). Wax from etiolated leaf sheaths consisted of 9% alkylresorcinols (0.3 ± 0.03 µg/cm2), 6% alkyl esters (0.2 ± 0.01 µg/cm2), 2% alkanes (0.1 ± 0.01 µg/cm2) and 2% fatty acids (0.1 ± 0.01 µg/cm2). Wax extracted from etiolated cotyledons comprised 11% fatty acids (0.4 ± 0.05 µg/cm2), 10% alkyl esters (0.3 ± 0.05 µg/cm2), 9% primary alcohols (0.3 ± 0.03 µg/cm2), and  Fatty acidsPrimary alcoholsAlkanesEstersAldehydesAlkylresorcinolsOthersWax coverage [µ g/cm2 ]024681012 Green leaf bladesEtiolated leaf bladesGreen external sheathsGreen internal sheathsEtiolated stemsEtiolated hypocotylsEtiolated cotyledonsEtiolated sheaths34  2% alkanes (≤ 0.1 µg/cm2). In contrast to green organs, aldehydes could not be detected in any of the etiolated organs.  3.2.1.4 Chain length distributions in waxes from etiolated and green S. cereale organs  Homologous series of fatty acids (C18-C34), primary alcohols (C18-C34), alkanes (C22-C29), aldehydes (C24-C38), alkyl esters (C38-C52) and alkylresorcinols (C19-C27) were identified in all S. cereale organs. However, the chain length distributions differed substantially, both among the green (Fig 3.4) and the etiolated (Fig 3.5) S. cereale organs. The free fatty acid fractions were dominated by the C26 homolog, amounting to 55% of the compound class (0.07 ± 0.01 µg/cm2) in wax on green leaf blades, 38% (0.04 ± 0.02 µg/cm2) on green external sheaths, 56% (0.2 ± 0.01 µg/cm2) on green internal sheaths, 25% (0.03 ± 0.004 µg/cm2) on etiolated leaf blades and 36% (0.1 ± 0.04 µg/cm2) on etiolated stems. In the waxes from etiolated hypocotyls and cotyledons, the C24 acid homolog was predominant with 66% (0.4 ± 0.06 µg/cm2), and 78% (0.2 ± 0.04 µg/cm2), respectively. Finally, etiolated sheaths had the C30 chain length as dominant fatty acid with 36% (0.02 ± 0.001 µg/cm2).          35                Figure 3.4 Chain length distributions within compound classes in waxes from green S. cereale organs. Relative amounts of homologs are given as % of the wax fraction from green leaf blades, external and internal sheaths. The percentage of each homolog within the compound class is given as mean value (n = 6) ± SD.  The primary alcohol fractions within waxes from all rye organs were dominated by even-numbered homologs, with hexacosanol being the most abundant homolog on green leaf blades with 94% (10.1 ± 1.8 µg/cm2), on green external sheaths with 89% (2.2 ± 0.6 µg/cm2), on green internal sheaths with 91% (2.0 ± 0.03 µg/cm2), on etiolated leaf blades with 66% (4.9 ± 1.4 µg/cm2), on etiolated sheaths with 44% (0.4 ± 0.06 µg/cm2), and on etiolated stems with 39% (0.7 ± 0.1 µg/cm2). The C22 alcohol homolog was predominant in waxes from ic  18ic  20ic  22ic  24ic  26ic  28ic  30ic  32ic  34ol  18ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  23an  26an  27an  28an  29an  30an  31an  32an  33 ..al  24al  26al  28al  30al  32al  34al  38020406080100Green leaf bladesic  18ic  20ic  22ic  24ic  26ic  28ic  30ic  32ic  34ol  18ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  23an  26an  27an  28an  29an  30an  31an  32an  33 ..al  24al  26al  28al  30al  32al  34al  38020406080 Green external sheathsic  18ic  20ic  22ic  24ic  26ic  28ic  30ic  32ic  34ol  18ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  23an  26an  27an  28an  29an  30an  31an  32an  33 ..al  24al  26al  28al  30al  32al  34al  38020406080 Green internal sheathsFatty acids Primary alcohols Alkanes AldehydesRelative composition [% of compound class]36  ic   18ic   20ic   22ic   24ic   26ic   28ic   30ic   32ic   34ol   18ol   20ol   21ol   22ol   23ol   24ol   25ol   26ol   27ol   28ol   30ol   32ol   34.an   22an   23an   26an   27an   2901020304050607080ic   18ic   20ic   22ic   24ic   26ic   28ic   30ic   32ic   34ol   18ol   20ol   21ol   22ol   23ol   24ol   25ol   26ol   27ol   28ol   30ol   32ol   34.an   22an   23an   26an   27an   2901020304050607080Etiolated leaf bladeic   18ic   20ic   22ic   24ic   26ic   28ic   30ic   32ic   34ol   18ol   20ol   21ol   22ol   23ol   24ol   25ol   26ol   27ol   28ol   30ol   32ol   34.an   22an   23an   26an   27an   29010203040506070 Etiolated stemic   18ic   20ic   22ic   24ic   26ic   28ic   30ic   32ic   34ol   18ol   20ol   21ol   22ol   23ol   24ol   25ol   26ol   27ol   28ol   30ol   32ol   34.an   22an   23an   26an   27an   29010203040506070 Etiolated sheathEtiolated hypocotylic   18ic   20ic   22ic   24ic   26ic   28ic   30ic   32ic   34ol   18ol   20ol   21ol   22ol   23ol   24ol   25ol   26ol   27ol   28ol   30ol   32ol   34.an   22an   23an   26an   27an   2901020304050607080 Etiolated cotyledonRelative composition [% of compound class]Fatty acids Primary alcohols AlkanesFatty acids Primary alcohols Alkanesetiolated hypocotyls and cotyledons, with 23% (0.2 ± 0.01 µg/cm2) and 33% (0.1 ± 0.05 µg/cm2), respectively. Moreover, all etiolated organs also contained higher amounts of C22 and C24 primary alcohols than green organs, comprising 16% and 12% on etiolated leaf blades, 30% and 15% on etiolated leaf sheaths, and 23% and 11% on etiolated stems, respectively.                 Figure 3.5 Chain length distributions within compound classes in waxes from etiolated S. cereale organs. Relative amounts of homologs are given as % of the fraction within waxes from leaf blades, stems, sheaths, hypocotyls and cotyledons. The percentage of each homolog within the compound class is given as mean value (n = 6) ± SD.  The wax alkanes from all green organs had chain length distributions dominated by the C29 homolog, at 65% (0.3 ± 0.08 µg/cm2) on green leaf blades, 50% (0.2 ± 0.04 µg/cm2) on green 37  external sheaths, 50% (0.1 ± 0.04 µg/cm2) on green internal sheaths, and at 59% (≤ 0.1 µg/cm2) on etiolated leaf blades. All other etiolated organs showed alkane profiles culminating at the C27 homolog, with 59% in etiolated sheaths, 45% in etiolated stems and hypocotyls, and 56% in cotyledons (all amounts ≤ 0.1 µg/cm2). Alkanes with chain length higher than C29 were not detected in etiolated organs.   Aldehydes with only even-numbered homologs were identified in waxes from green organs. C26 represented the predominant chain length, with approximately 91% (1.7 ± 0.5 µg/cm2) on green leaf blades, 86% (1.0 ± 0.1 µg/cm2) on green external sheaths and 95% (0.6 ± 0.05 µg/cm2) on green internal sheaths.    3.2.2 Cuticular wax profile of various Brachypodium distachyon organs. Total wax mixtures were extracted from entire B. distachyon organs, under the same conditions as described for S. cereale (see 3.2.1). The total wax amounts on various Brachypodium distachyon organs were quantified (3.2.2.1), together with compound class distributions (3.2.2.2) and their respective chain length profiles (3.2.2.3).  3.2.2.1 Total wax amount of Brachypodium distachyon organs Waxes were extracted with chloroform from six green Brachypodium distachyon organs (cotyledons, spikes, leaf blades, sheaths, stems and internal stems), then transformed into trimethylsilyl (TMSi) derivatives and analyzed on GC-MS and GC-FID. Cotyledons (14.1 ± 1.9 µg/cm2), spikes (13.8 ± 1.1 µg/cm2) and leaf blades (13.3 ± 1.2 µg/cm2) had similarly 38  high wax coverages (Fig 3.6), in contrast to lower levels on sheaths (5.1 ± 0.1 µg/cm2), exposed stems (3.0 ± 0.4 µg/cm2) and covered stem parts (2.4 ± 0.5 µg/cm2).  CotyledonsSpikesLeaf bladesSheathsStemsInternal stemsWax coverage [µ g/cm2 ]024681012141618 Figure 3.6 Total wax loads on various Brachypodium distachyon organs. Wax coverages are given as mean values (n = 6) ± SD.  3.2.2.2 Compound class distribution in waxes on Brachypodium distachyon organs Waxes from various Brachypodium distachyon organs comprised typical compound classes such as fatty acids, primary alcohols, alkanes, aldehydes and alkyl esters, together with alkylresorcinols (Fig 3.7). The relative amounts of these compound classes differed between organs, however primary alcohols were the predominant compound class throughout. For instance, wax on cotyledons contained 56% (7.8 ± 0.6 µg/cm2), on spikes 53% (7.2 ± 0.8 µg/cm2), on leaf blades 57% (7.6 ± 0.8 µg/cm2), on sheaths 45% (2.2 ± 0.3 µg/cm2), on stems 49% (1.5 ± 0.2 µg/cm2), and on internal stems 52% (1.1 ± 0.3 µg/cm2) of primary alcohols. 39           Figure 3.7 Compound class composition of wax mixtures from various B. distachyon organs. Waxes from cotyledons, spikes, leaf blades, leaf sheaths, stems and internal stems were analyzed. The coverage of each compound class is given as mean value in µg/cm2 ± SD.   Alkyl esters were the second major compound class on leaf blades with ca 13% (1.7 ± 0.2 µg/cm2), on spikes with ca 26% (2.2 ± 0.2 µg/cm2), on cotyledons with ca 10% (1.3 ± 0.1 µg/cm2), and on internal stems with ca 6% (0.2 ± 0.04 µg/cm2). Waxes on leaf sheaths and stems contained ca 18% of aldehydes, on leaf blades ca 8% (1.1 ± 0.1 µg/cm2), on spikes ca 15% (2.1 ± 0.2 µg/cm2), on cotyledons ca 6% (0.7 ± 0.1 µg/cm2), and on internal stems ca 4% (0.2 ± 0.02 µg/cm2). Alkylresorcinols were found only on leaf blades at ca 6% (0.8 ± 0.1 µg/cm2), and on leaf sheaths at ca 1% of the total wax. Finally, alkanes and fatty acids contributed minor percentages on all organs, with concentrations of 2% and 1% on leaf blades, 2% and 1% on spikes, 4% and 1% on cotyledons, 4% and 1% on internal stems, 1% and 2% on leaf sheaths, and 2% and 6% on stems, respectively.  Fatty acidsAlcoholsAlkanesAlkyl estersAldehydesAlkylresorcinolsOthersWax coverage [µ g/cm2 ]0246810CotyledonsSpikesLeaf bladesSheathsStemsInternal stems40  3.2.2.3 Chain length distribution in waxes from Brachypodium distachyon organs Homologous series of fatty acids (C18-C30), primary alcohols (C20-C34), alkanes (C22-C33), aldehydes (C24-C34), alkyl esters (C38-C54) and alkylresorcinols (C17-C25) were present in waxes from all Brachypodium distachyon organs (Fig 3.8). However, the chain length distributions differed among Brachypodium distachyon organs. Free fatty acids were dominated by the C26 homolog, amounting to 39% of the compound class on cotyledons, 33% on spikes, 60% on leaf blades, and 39% on leaf sheaths. In the waxes from stems and internal stems, the C28 fatty acid homolog was predominant, with 84%, and 42%, respectively. Overall, the major fatty acid homologs had concentrations of less than 0.1 µg/cm2 in all organs, except on stems with a C28 acid coverage of 0.1 ± 0.02 µg/cm2.   The primary alcohol fractions in the waxes from all Brachypodium distachyon organs were dominated by even-numbered homologs ranging from C20 to C34 carbons in length, with C26 as the predominant homolog throughout. The alcohol fraction on cotyledons comprised 96% (7.5 ± 0.6 µg/cm2) of the C26 homolog, compared to 92% (6.7 ± 0.9 µg/cm2) on spikes , 92% (7.0 ± 0.8 µg/cm2) on leaf blades, 87% (1.9 ± 0.2 µg/cm2) on leaf sheaths, 83% (1.2 ± 0.2 µg/cm2) on stems, and 94% (1.0 ± 0.2 µg/cm2) on internal stems.   The wax alkane chain length distribution was dominated by the C29 homolog on leaf blades with 35% (0.1 ± 0.02 µg/cm2), and on spikes with 51% (0.2 ± 0.02µg/cm2). In contrast, C31 alkane was dominant on cotyledons with 44% (0.3 ± 0.02 µg/cm2), on stems with 46% (≤ 0.1 µg/cm2), and on internal stems with 63% (≤ 0.1 µg/cm2). Finally, the C27 homolog was predominant on sheaths with 66% (≤ 0.1 µg/cm2).  41  Within the aldehyde fractions, only even-numbered homologs were identified. C26 aldehyde dominated throughout, on cotyledons with 84% (0.7 ± 0.1 µg/cm2), on spikes with 66% (1.4 ± 0.4 µg/cm2), on leaf blades with 74% (0.8 ± 0.05 µg/cm2), on leaf sheaths with 63% (0.6 ± 0.1 µg/cm2), on stems with 65% (0.4 ± 0.04 µg/cm2), and on internal stems with 66% (0.1 ± 0.01 µg/cm2).              Figure 3.8 Chain length distributions within compound classes in waxes from various Brachypodium distachyon organs. Relative amounts of homologs are given as mean percentages of the fraction within waxes from cotyledons, spikes, leaf blades, sheaths, stems, and internal stems (n = 6) ± SD. ic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080100Cotyledonsic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080ic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080ic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080100ic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080ic  18ic  20ic  22ic  24ic  26ic  28ic  30ol  20ol  21ol  22ol  23ol  24ol  25ol  26ol  27ol  28ol  30ol  32ol  34.an  22an  27an  28an  29an  30an  31an  33 ..al  24al  26al  28al  30al  32al  34020406080Relative composition [% of compound class]SpikesLeaf bladesSheathsStemsInternal stemsFatty acids Primary alcohols Alkanes Aldehydes Fatty acids Primary alcohols Alkanes Aldehydes42  3.3 Discussion The main objective in this chapter was to analyze the components of cuticular waxes on various organs of Brachypodium distachyon and S. cereale grown in the light or dark. Details of the amounts of alkylresorcinols and alkyl esters contained in the wax mixtures will be discussed in chapters 4 and 5, respectively  3.3.1 Total wax of green and etiolated S. cereale organs S. cereale leaf blades were found to have higher wax amounts than the other organs investigated in this study. This finding confirmed values reported previously for leaf wax of S. cereale (Ji & Jetter, 2008; Streibl et al., 1974; Tulloch & Hoffman, 1974) and other related grass species (Tulloch, 1981). Furthermore, wax composition is known to vary between different organs (Buschhaus & Jetter, 2011; Jenks et al., 1995;  Jetter & Schäffer, 2001; Rashotte et al., 1997; Shepherd & Wynne Griffiths, 2006) and drastic differences have  been reported between cuticular wax compositions of various grass organs, for instance on barley leaves and spikes (Simpson & Wettstein-Knowles, 1980; Wettstein-Knowles & Netting, 1976), on Triticum aestivum spikes, leaf blades and sheaths (Tulloch, 1973; Tulloch & Hoffman, 1973; Bianchi et al., 1979). Similarly differing wax coverages were found here for the various organs of S. cereale selected.  All the green organs had relatively high amounts of waxes compared with the etiolated tissues. In line with this result, also the waxes on green internal leaf sheaths, which are exposed to only little light penetrating the surrounding tissues, had lower coverages compared with those on external sheaths. The total wax coverages on etiolated leaf blades 43  decreased to ca 75% compared with the same organ in the light, and etiolated sheaths had ca 43% of the wax on green sheaths. Overall, this study shows that cuticular wax formation and/or deposition are affected by light levels during organ growth. This finding matches previous reports on Allium porrum L. (leek), where etiolated seedlings had 80% of those grown in the light (Maier & Post-Beittenmiller, 1998), and on Hordeum vulgare (Bonus barley), where etiolated leaves had ca 60% of the wax than those grown in the light (Giese, 1975). Conversely, waxes on B. oleracea  (Baker, 1974) and Brassica napus (Whitecross & Armstrong, 1972) leaves had been shown to accumulate to higher amounts when grown in high-light conditions. Therefore, the relatively low wax levels found in the present study for etiolated S. cereale organs confirms previous reports on the effect of light on wax biosynthesis/deposition.  3.3.2 Compound class and homolog distributions in cuticular waxes from S. cereale organs Not only total wax amounts but also compositions are known to vary among species, organs, tissues and developmental stages. In Arabidopsis thaliana, differences of cuticular wax composition have been reported between stems (Hannoufa et al., 1993), leaves (Jenks et al., 1995), pollen (Preuss et al., 1993) and siliques (Koornneef et al., 1989). Similarly, wax compositions also differed between organs of grass species, for example between barley spikes and leaf blades (Simpson & Wettstein-Knowles, 1980), between wheat spikes, leaf blades and sheaths (Bianchi et al., 1979), and between Zea mays leaves and pollen (Bianchi et al., 1990). Overall, previous reports had shown that most grass species and organs, including S. cereale leaves, had waxes dominated by primary alcohols (Ji & Jetter, 2008; Tulloch & Hoffman, 1974). Accordingly, the wax compositions found here for S. cereale 44  organs were similar to those reported in other Poaceae species such as Oryza sativa (Bianchi et al., 1979), T. aestivum (Tulloch & Hoffman, 1974),  H. vulgare (Reynhardt and Riederer, 1994) and Z. mays (Żarnowski et al., 2002; Ross et al., 2004).   Hexacosanol was the predominant alcohol homolog (except for etiolated hypocotyls and etiolated cotyledons), confirming previous findings from a study of 174 species of the tribe Triticeae (Tulloch, 1981). The predominant C26 primary alcohol had also been reported before for leaf waxes from H. vulgare (Richardson et al., 2005) and S. cereale (Ji & Jetter, 2008). However, in T. aestivum leaves (Koch et al., 2006) the C28 homolog had been found be prevalent, a predominance of the C30 alcohol homolog had been reported for leaves of O. sativa (Yu et al., 2008), and the C32 homolog dominated on seedlings of Z. mays (Bianchi et al., 1989).   In the current research, etiolated leaf blades had a decrease in C26 primary alcohol and an increase of C22 and C24 alcohols compared with green leaf blades (Fig 3.9). Etiolated sheaths also presented the same variation between C22, C24 and C26, with decreased levels of C26 and increased levels of C22 and C24. Similar tendencies had previously been reported for H. vulgare leaves (Giese, 1975), demonstrating that etiolation results in a broader homolog distribution.      45                     Figure 3.9 Comparison of chain length distributions of individual wax components in green and etiolated leaf blades and sheaths. Percentages of individual homologs within the series of (A) fatty acids, (B) primary alcohols and (C) alkanes in green and etiolated leaf blades, and of (D) fatty acids, (E) primary alcohols and (F) alkanes in green and etiolated leaf sheaths are shown as mean values (n = 6) ± SD.  Fatty acids chain length18 20 22 24 26 28 30 32 340102030405060Green leaf blades Etiolated leaf blades Alcohols chain length18 20 21 22 23 24 25 26 27 28 30 32 34Relative composition [%]020406080100Alkanes chain length22 27 28 29 30 31 32 33020406080Fatty acids chain length18 20 22 24 26 28 30 32 340102030405060Green external sheaths Green internal sheathsEtiolated sheaths Alcohols chain length18 20 21 22 23 24 25 26 27 28 30 32 34020406080100Alkanes chain length22 27 28 29 30 31 32 33020406080ABCDEF46  Aldehydes were not detected in etiolated S. cereale organs, similar to findings for H. vulgare (Giese, 1975) and Brassica. oleracea (Macey, 1970). In all green tissues investigated here, the predominant homolog was C26 aldehyde, confirming values reported previously for S. cereale leaves (Ji & Jetter, 2008). However, aldehydes had not been reported by Tulloch and Hoffman (1974). It should be noted that aldehydes and primary alcohols shared similar homolog distributions in all S. cereale organs, matching previous reports on other grass species and leading to the notion that both compound classes may be biosynthetically related. Accordingly, it has been postulated that the grass wax aldehydes may be intermediates in route to primary alcohols (Jetter & Kunst, 2008), in contrast to the situation in Arabidopsis thaliana, where aldehydes that are occur as intermediates in the biosynthesis of wax alkanes instead.  Alkanes were detected in all S. cereale organs, when grown in the light with C29 as predominant homolog. For S. cereale leaves, a similar alkane composition had been reported once (Tulloch & Hoffman, 1974), however another study had also shown C31 alkane as the predominant compound (Ji & Jetter, 2008). On etiolated organs, the number of wax alkane homologs was reduced considerably. In etiolated leaf blades, only C29 (55%) and C27 (45%) were found. In contrast, the other etiolated organs had broader distributions with C27 as predominant alkane chain length, thus leading to a shift to shorter chains than on green organs.   Fatty acids were found at relatively high concentrations in waxes at early developmental stages such as etiolated cotyledons and etiolated hypocotyls, in sharp contrast to all late-47  developing organs with only low levels of free fatty acids. This finding indicates that drastically different wax biosynthetic mechanisms may be operating during early stages of development, accentuated in the dark. Developmental changes in wax production had been observed before in Z. mays and Coix lachrymal (Avato et al., 1990) and Kalanchoe daigremontiana (Van Maarseveen et al., 2009).   Here, the C26 fatty acid homolog was predominant in all green S. cereale organs, thus matching the chain length profiles of alcohols and aldehydes in respective wax mixtures. The waxes from etiolated organs were found to have more varied fatty acid chain length patterns, where early-developing organs had shorter fatty acid homologs than those from later developmental stages.   Overall, this study showed clear differences between homolog distributions in green and etiolated organs. In the light, homolog patterns were dominated by C29 alkane and C26 primary alcohol, aldehyde and fatty acid, while in the dark these compound classes showed more even chain length distributions, suggesting that light affects the biosynthesis and/or deposition of cuticular waxes. It is possible that cuticular wax compositions depend on photoperiods, as de novo fatty acid synthesis in chloroplasts increases in the light. The synthesis of malonyl-CoA, catalyzed by acetyl-CoA carboxylase (ACCase), is the first committed step in fatty acid biosynthesis, and it is modulated by light-dark conditions (Sasaki et al., 1997)   48  3.3.4 Total wax of Brachypodium distachyon organs Brachypodium distachyon has emerged as a relatively new model system, where no chemical analyses in waxes from different organs had been reported yet. To understand the wax distribution between different organs, analyses of waxes were performed in the current study. Etiolated Brachypodium distachyon organs could not be analyzed as specimens could not be grown reproducibly in the dark.  Leaf blades had a total wax coverage of 13.3 ± 1.2 µg/cm2, similar to previously reported values (Yao, 2011). The total wax levels on cotyledons, on spikes and on leaf blades were relatively similar to those on leaf blades, while the other organs had lower wax amounts. This finding again confirms previous reports on drastic variations between wax coverages on different organs. For instance, tomato fruit had a wax coverage of 15 µg/cm2, contrasting with only 3 µg/cm2 on leaves (Vogg et al., 2004). It is currently not clear how the total amount of waxes may influence their physiological and ecological role, except for the notion that high wax amounts may be a prerequisite for the formation of surface wax crystals (Simpson & Wettstein-Knowles, 1980; Wettstein-Knowles & Netting, 1976) that are important in modulating herbivore behaviour (Riederer & Muller, 2006)  3.3.5 Homolog distributions in cuticular waxes from Brachypodium distachyon organs  The waxes from all Brachypodium distachyon organs selected in this research were dominated by primary alcohols and the homolog C26 in particular.  this finding matches numerous reports on other Gramineae, for example on H. vulgare (Richardson et al., 2005) and on S. cereale (Ji & Jetter, 2008). The C26 homolog was also predominant in the fatty acid 49  and aldehyde fractions from the majority of Brachypodium distachyon organs. Similar to S. cereale, aldehydes in Brachypodium distachyon therefore seem to be biosynthetically related to alcohols and acids, most likely as intermediates in the reduction pathway leading from acyl CoAs to alcohols (Jetter & Kunst, 2008).   The wax alkanes from various Brachypodium distachyon organs had broad homolog distributions. For instance, C29 was the predominant homolog on spikes and leaf blades, C31 on cotyledons,   and stems, and C27 on leaf sheaths. Similar alkane chain length variability had been reported before, indicating a broad distribution between species with major percentages of C27, C29, C31 and C33 in Gramineae (Tulloch, 1981)              50  Chapter 4: Cuticular alkylresorcinol homologs in various organs of Secale cereale and Brachypodium distachyon 4.1 Introduction 5-Alkylresorcinols (ARs) are a class of bioactive, polyketide-derived phenolic lipids, also called 1,3-dihydroxy-5-alkylbenzenes or 5-n-alkylresorcinols. Their amphiphilic properties are due to a combination of a hydrophobic alkyl chain and a resorcinol ring. Many derivatives of alkylresorcinols have been identified in a large variety of plants and animals (Kozubek & Tyman, 1999). For instance, in cereal species ARs had been detected in non-edible organs such as seedlings of Oryza sativa, Secale cereale and Zea mays (Deszcz & Kozubek, 2000; Suzuki et al. , 1996; Suzuki et al, 2003), roots of Sorgum bicolor (Cook et al., 2010), and leaves of Secale cereale (Ji & Jetter, 2008).  In addition, ARs had been localized in the bran fraction of whole grain of wheat and rye (Chen et al., 2004), in the milling fraction of wheat, rye and bread (A. B. Ross et al., 2003), in the latex and peel of Mangifera indica (Droby et al., 1986) and in seed covers of Mangifera fragrans and shells of Anacardium occidentale (cashew nut) (Kozubek & Tyman, 1999). Overall, these occurrences suggested that the ARs are deposited on plant surfaces, or more specifically, in the cuticles of various organs and species. Accordingly, ARs were found in the cuticular wax of Hordeum vulgare grains (García et al., 1997), and a recent study of Secale cereale confirmed the accumulation of alkylresorcinols in leaf cuticles (Ji & Jetter, 2008), more specifically localizing ARs in the intracuticular wax layer.  The composition of very-long-chain alkylresorcinol in leaf waxes of Secale cereale were investigated, indicating that AR homologs occur in relatively high levels when the second 51  leaves of plants are 20 cm long (15 - 17 days old) (Ji & Jetter, 2008). It was also reported that cuticular wax on B. distachyon leaves contains substantial amounts of ARs, albeit with slightly shorter chain lengths than rye leaves (Yao, 2011). The homologous series of alkylresorcinols in leaf waxes of Secale cereale and B. distachyon were identified using GC-MS (Ji & Jetter, 2008; Yao, 2011). TMSi derivatives of all the alkylresorcinols shared characteristic MS fragments m/z 73, 268, 281 (Linko et al., 2002; Ross et al., 2003). All the homologs had different molecular ions [C6H3(OTMS)2(CH2)nCH3]+ accompanied by fragments [M-15]+ due to loss of a methyl group. The pairs of these two ions differed by m/z 28 between homologs, thus confirming that all respective compounds belonged to a series differing by -CH2CH2- units. The homologous series of alkylresorcinols identified in S. cereale and B. distachyon leaves had side chains with odd numbers of carbons ranging from C19 to C27 (Ji & Jetter, 2008), and C17 to C25 (Yao, 2011), respectively.   A previous study identified two new ARs in waxes from B. distachyon leaves (Yao, 2011) as methyl alkylresorcinols C19 and C21. Similar compounds were also described once in wheat with chain lengths ranging from C19 to C27 (Adamski et al., 2013). In both reports, the methyl alkylresorcinol structures were inferred from the MS fragment m/z 282, but without assigning exact isomer structures. Three possible methyl positions were predicted: in the resorcinol ring in the ortho or para positions (RMARs), or at the 1’ position (aka as the benzylic position) on the alkyl side chain (CMAR) (Fig 4.1)    52       Figure 4.1 Possible positions of the methyl group in the new alkylresorcinol series. The methyl group may be located (A) on the resorcinol ring in the ortho position, (B) on the resorcinol ring in the para position, or else (C) in the (benzylic) position 1’ on the alkyl side chain.  The previous chemical identification of cuticular ARs in both Secale cereale and B. distachyon leaves suggested that one or more alkylresorcinol synthases (ARSs) produce alkylresorcinols for functions in the waxes, using VLCFA as starter units via condensations with malonyl-CoA. Two genes potentially involved in AR biosynthesis were isolated from Secale cereale (ScARS) and B. distachyon (BdARS) and partially characterized (Yao, 2011). In B. distachyon, gene cloning was performed based on the mining of B. distachyon expressed sequence tag (EST) libraries. In S. cereale, PCR-based gene cloning was carried out using primer designs exploiting homology between the functionally characterized ARS sequences from Sorghum bicolor and O. sativa (Yao, 2011). Yeast in vivo characterization showed that the corresponding proteins had ARS activity, however the AR homologs formed had shorter chains than those in the waxes of both plant species. It was therefore not clear whether the ARSs are indeed involved in formation of the cuticular ARs. In order to further test involvement of the genes/proteins in formation of the wax ARs, they had to be characterized. In particular, gene activity should be correlated with AR product accumulation. Therefore, comprehensive expression profiling in various organs and growth 282 282 282 A B C 53  conditions was to be performed in conjunction with wax profiling, and protein subcellular localization to assess access of the enzymes to substrate pools.  The goal of the current chapter was to identify and quantify the cuticular alkylresorcinols homologs in waxes on various organs Secale cereale and B. distachyon. To this end, also the new group of cuticular methyl alkylresorcinol homologs had to be identified using synthetic standards for comparison of GC-MS retention times and fragmentation patterns. Finally, further characterizations of ScARS and BdARS were performed, using gene expression profiling in different plant organs and subcellular localization, to enable correlations between gene activity and product accumulation.               54  4.2 Results The overall objective of this study is to analyze the alkylresorcinol homologs in the cuticular waxes of Secale cereale and B. distachyon. Further to previous studies reporting alkylresorcinol compositions for the leaf waxes of these two species (Ji & Jetter, 2008; Yao, 2011), the current research aimed at broadening our knowledge by first identifying all alkylresorcinols in the waxes of various Secale cereale and B. distachyon organs (4.2.1).  A principal goal of the present chapter was then to quantify all the alkylresorcinols within the complex wax mixtures on various organs of both species, and to compare compositions between tissues grown in light or in the dark (4.2.2). Finally, the previously isolated ScARS and BdARS genes had to be characterized by comparing expression levels across organs (4.2.3), and by determining the subcellular localization of their enzyme products (4.2.4). Taken together, correlations between the chemical and molecular datasets can help corroborate the involvement of the candidate genes in the biosynthesis of cuticular alkylresorcinols.   4.2.1 Identification of cuticular alkylresorcinols and methyl alkylresorcinols  To give a full account of alkylresorcinols in Secale cereale and B. distachyon waxes, first the homologs previously described for leaf waxes of both species had to be confirmed, additional homologs had to be identified, and corresponding series had to be detected in the waxes on other organs of both species (4.2.1.1). Second, a group of alkylresorcinol isomers, recently reported for B. distachyon leaf wax, had to be further identified and detected in various Secale cereale organs (4.2.1.2).   55   4.2.1.1 Identification of alkylresorcinol homologs in waxes from Secale cereale and B. distachyon Alkylresorcinols with side chains ranging from C19 to C27 in Secale cereale and C17 to C25 in B. distachyon leaf waxes had been identified before (Ji & Jetter, 2008; Yao, 2011) and their presence in the current leaf wax samples had to be confirmed first. To this end, GC-MS with selected ion monitoring was used to analyze the TMSi-derivatized wax mixtures. The co-occurrence of characteristic MS fragments m/z 268 (Fig. 4.2), as well as m/z 73 and 281 (data not shown) in GC peaks with equal distances showed the homologous nature of all compounds involved.   To confirm the alkylresorcinol structure for all the peaks in the series, their MS fragmentation patterns were analysed. The resulting MS fragmentation patterns matched those previously described for alkylresorcinols, including ions m/z 73 indicative of TMSi groups, m/z 268 and 281 characteristic of resorcinols, and molecular ions [C6H3(OTMSi)2(CH2)nCH3]+ accompanied by fragments [M-15]+ (Fig 4.3).         56                          Figure 4.2 GC-MS single ion chromatograms (m/z 268) of TMSi-derivatized leaf blade waxes of Secale cereale and B. distachyon. (A) Alkylresorcinols with C19 - C27 side chains were identified in wax from green leaf blades of Secale cereale, and (B) with side chains of C17 - C25 for B. distachyon leaf blades.       15 20 25 30 35 40 45Abundance02000040000600008000010000012000014000016000019:021:0 23:025:027:0ARetention time [min]15 20 25 30 35Abundance02000040000600008000010000012000014000016000019:021:023:025:017:0B57                        Figure 4.3 Mass spectra of TMSi-derivatized alkylresorcinols in wax of Secale cereale green leaf blades. (A) Alkylresorcinol with C19 side chain (AR 19:0; molecular ion m/z 520). (B) Alkylresorcinol with C21 side chain (AR 21:0; molecular ion m/z 548). (C) Alkylresorcinol with C23 side chain (AR 23:0; molecular ion m/z 576). (D) Alkylresorcinol with C25 side chain (AR25:0; molecular ion m/z 604). (E) Alkylresorcinol with C27 side chain (AR 25:0; molecular ion m/z 632). 020406080100020406080020406080020406080m/z50 100 150 200 250 300 350 400 450 500 550 600 650Relative Abundance [%]020406080268268268268268281281281281281 505520533548561576589604617632Alkylresorcinol 19:0Alkylresorcinol 21:0Alkylresorcinol 23:0Alkylresorcinol 25:0Alkylresorcinol 27:0ABCDE520 281 268 58  Finally, to confirm the exact structure, a standard of 5-n-nonadecylresorcinol (synthesized by former student X. Ji) was analyzed. It was found to have an MS fragmentation pattern and GC retention behaviour identical with one of the alkylresorcinol homologs (AR 19:0) in Secale cereale green leaf blades wax (Fig 4.4).                 Figure 4.4 MS analysis of TMSi-derivatized C19:0 alkylresorcinols. (A) Mass spectrum of alkylresorcinol with C19 side chain (AR 19:0; molecular ion m/z 520) in wax from Secale cereale green leaf blades, (B) synthetic standard of C19:0 alkylresorcinol, and (C) structure and fragmentation pattern of 5-n-nonadecylresorcinol.  In wax of green Secale cereale leaf blades, the alkylresorcinols with odd-numbered alkyl chain lengths ranging from C19 to C27 were identified (Fig.4.2). In addition, homologs with even-numbered side chains of C20, C22 and C24 were also detected. The alkylresorcinol 020406080100268281 505520Alkylresorcinol 19:0m/z50 100 150 200 250 300 350 400 450 500 550 600 650020406080281268505520Standard alkylresorcinol 19:0ABRelative Abundance [%]C   59  homologs found in all other (green and etiolated) organs of Secale cereale had the same chain lengths as in green leaf blades (Table 4.1), except in etiolated cotyledons, where no alkylresorcinols could be detected. In addition traces of ARs homolog C29 and C31 were detected in etiolated hypocotyl, and a series of unsaturated ARs odd-numbered C19 to C27 were detected in etiolated leaf blades and etiolated sheaths (data not shown). Only in leaf blades and sheaths of B. distachyon could alkylresorcinols be detected, and there the series included odd-numbered alkyl chain length ranged from C17 to C25, and even-numbered C20 and C22.  Table 4.1 List of alkylresorcinol homologs identified in various S. cereale and B. distachyon organs.              tr traces; nd no detected      Alkylresorcinol homologs    C17 C19 C20 C21 C22 C23 C24 C25 C27 Secale cereale Green organs           Leaf blades nd ✓ tr ✓ tr ✓ tr ✓ ✓ External sheaths nd ✓ tr ✓ tr ✓ tr ✓ ✓ Internal sheaths nd ✓ tr ✓ tr ✓ tr ✓ ✓            Etiolated organs Leaf blades nd ✓ tr ✓ tr ✓ tr ✓ ✓ Sheaths nd ✓ tr ✓ tr ✓ tr ✓ ✓ Stems nd ✓ tr ✓ tr ✓ tr ✓ ✓ Hypocotyls nd ✓ tr ✓ tr ✓ tr ✓ ✓ Cotyledons nd nd  nd nd nd nd nd nd nd             Brachypodium distachyon  Leaf blades ✓ ✓ tr ✓ tr ✓ nd ✓ nd  Sheaths ✓ ✓ nd ✓ nd ✓ nd nd nd  Stems nd nd nd nd nd nd nd nd nd  Internal stems nd nd nd nd nd nd  nd nd nd  Spikes nd nd nd nd nd nd nd nd nd  Cotyledons nd nd nd nd nd nd nd nd nd 60  4.2.1.2 Identification of methyl alkylresorcinols in waxes from S. cereale and B. distachyon Two new compounds in B. distachyon leaf wax had been suggested to be isomeric C19 and C21 alkylresorcinols based on characteristic fragments m/z 282 (Yao, 2011) . In order to further elucidate the structure of these compounds and to confirm their presence also in Secale cereale waxes, detailed MS analyses were carried out. The new compounds were indeed detected in the wax mixture on green Secale cereale leaf blades, where they formed a homologous series defined by similar MS fragmentation patterns and equal distances between GC peaks (Fig 4.7-A). All compounds were baseline-separated from normal alkylresorcinols, showing a pronounced pattern of high abundance for every other homolog. The abundant homologs eluted between alkylresorcinols with odd and even chain lengths, whereas the less common homologs eluted between even- and odd-numbered alkylresorcinols.  All the novel compounds had characteristic MS fragments m/z 282 and m/z 295 (Fig 4.5), differing from the corresponding ions of normal alkylresorcinols by 14 additional mass units. The molecular ions [C6H3(OTMSi)2(CH2)nCH3]+ and fragments [M-15]+ were also similar to the normal alkylresorcinols, thus indicating isomeric structures. All the evidence taken together, the new structures were recognized as alkylresorcinols with an additional methyl group either at the 1’ position of the side chain or in the ortho or para positions on the ring, as previously hypothesized (Yao, 2011). Each methyl alkylresorcinol homolog eluted after the corresponding normal alkylresorcinol with the same chain lengths but 14 Da lower molecular mass, and before the normal alkylresorcinol with one carbon longer chain and the same molecular mass. 61                   Figure 4.5 Mass spectra of TMSi derivatives of methyl alkylresorcinols (MARs) in wax of green Secale cereale leaf blades. (A) Methyl alkylresorcinol C19 (MAR 19:0; molecular ion m/z 534). (B) Methyl alkylresorcinol C21 (MAR 21:0; molecular ion m/z 562). (C) Methyl alkylresorcinol C23 (MAR 23:0; molecular ion m/z 590). (D) Methyl alkylresorcinol C25 (MAR 25:0; molecular ion m/z 618).  Methyl alkylresorcinols with side chains ranging from C19 to C25 were detected (Fig 4.7A) in wax of green Secale cereale leaf blades. The methyl alkylresorcinol homologs found in all other (green and etiolated) organs of Secale cereale had the same chain lengths as in green 02040608010002040608002040608050 100 150 200 250 300 350 400 450 500 550 600 650Relative Abundance [%]020406080282519534547562575590603618Methyl alkylresorcinol 19:0Methyl alkylresorcinol 21:0Methyl alkylresorcinol 23:0Methyl alkylresorcinol 25:0m/z282282282ABCD B  D 62  leaf blades (Table 4.2). In etiolated cotyledons, no methyl alkylresorcinols could be detected. Only in leaf blades of B. distachyon, two methyl alkylresorcinols could be detected, C19 and C21 chain length.   Table 4.2 List of methyl alkylresorcinol homologs identified in various S. cereale and B. distachyon organs.                To further distinguish between the three remaining isomeric structures (Fig 4.1), methyl alkylresorcinol standards had to be synthesized for comparisons of GC-MS characteristics. It was speculated that the three isomers would show only minor differences in their MS    Methyl alkylresorcinol homologs    C19 C21 C23 C25 Secale cereale Green organs      Leaf blades ✓ ✓ ✓ ✓ External sheaths ✓ ✓ ✓ ✓ Internal sheaths ✓ ✓ ✓ ✓       Etiolated organs Leaf blades ✓ ✓ ✓ ✓ Sheaths ✓ ✓ ✓ ✓ Stems ✓ ✓ ✓ ✓ Hypocotyls ✓ ✓ ✓ ✓ Cotyledons nd nd  nd nd        Brachypodium distachyon  Leaf blades ✓ ✓ nd nd  Sheaths nd nd nd nd  Stems nd nd nd nd  Internal stems nd nd nd nd  Spikes nd nd nd nd  Cotyledons nd nd nd nd tr traces; nd no detected 63  fragmentation patterns, but would have different GC retention times and should thus be separable. However, the only synthetic standard at the time of this research (prepared by UBC Chemistry graduate student Chen Peng) was a methyl alkylresorcinol with the methyl group in the 1’ position on the alkyl side chain (CMAR).  The MS fragmentation pattern of CMAR with a side chain length of C21 was found to be very similar to that of C21 normal alkylresorcinol, except for subtle differences in the relative abundance of fragments m/z 57, 73, 267 and 323  (Fig 4.6).            Figure 4.6 Comparison of MS fragmentation patterns between methyl alkylresorcinols from Secale cereale wax and organic synthesis. (A) Mass spectrum of C21 methyl alkylresorcinol (MAR 21:0; molecular ion m/z 562) in Secale cereale green blade, and (B) mass spectrum of C21 methyl alkylresorcinol synthetic standard (CMAR 21:0; molecular ion m/z 562).  To test whether the methyl alkylresorcinols contained in the Secale cereale leaf blades wax and the synthetic CMAR standard could be distinguished by GC, both samples were analyzed 020406080100m/z50 100 150 200 250 300 350 400 450 500 550 600 650020406080547562MAR 21:0282547562CMAR 21:0282Relative Abundance [%]AB64  under identical chromatographic conditions and in the presence of C24 alkane standard. The retention times of the C24 alkane standard in both GC runs was very similar, at 11.66 min and at 11.62 min when measuring at the onset of the GC peak to avoid possible bias from varying concentrations (Fig 4.7). In contrast, the C21 methyl alkylresorcinol (MAR) in the wax sample from green leaf blades eluted much later (at 28.37 min) than the C21 methyl alkylresorcinol synthetic standard (CMAR) (at 26.81 min).                  Figure 4.7 Comparison of GC retention times between methyl alkylresorcinols from Secale cereale wax and organic synthesis. (A) Secale cereale green leaf blades sample showing four methyl alkylresorcinol (MAR) homologs C19, C21, C23 and C25, and C24 alkane internal standard. (B) Synthetic methyl alkylresorcinol (CMAR) with methylation in the 1’ position on the alkyl side chain, and C24 internal standard. The black trace (m/z 268) shows the (TMSi-derivatized) C19 to C27 alkylresorcinols, the red trace (m/z 282) the C19 to C27 methyl alkylresorcinols. 10 15 20 25 30 35 40Abundance0200040006000800010000120001400016000AR19:0AR21:0AR23:0AR25:0AR27:0MAR 19:0MAR 21:0MAR 23:0 MAR 25:0Retention time [min]10 15 20 25 30 35 40010002000300040005000CMAR 21:0AR22:024 alkane24 alkaneAR13:0AB11.62 min 11.66 min 28.37 min 26.81 min 65  4.2.2 Quantification of alkylresorcinols in waxes from various Secale cereale and B. distachyon organs In order to quantify all alkylresorcinol and methyl alkylresorcinol homologs identified in 4.2.1, corresponding peaks in GC-FID runs of total wax mixtures were assigned based on retention times and in comparison with select GC-MS runs. Synthetic alkylresorcinol 13:0 was co-injected as an internal standard together with the wax samples, and all individual alkylresorcinols (4.2.2.1) and methyl alkylresorcinols (4.2.2.2) were quantified by integrating their peak areas against that of the standard. Total amounts of alkylresorcinols in each wax mixture were determined by adding all alkylresorcinol and methyl alkylresorcinol homolog quantities together (4.2.2.3).   4.2.2.1 Alkylresorcinol chain length distribution in Secale cereale and B. distachyon organs Normal alkylresorcinol homologs were quantified where present in Secale cereale and B. distachyon organs, and expressed as percent of the total alkylresorcinol amounts. The waxes on all green Secale cereale organs (leaf blades, external and internal leaf sheaths) contained predominantly C23 alkylresorcinol, together with substantial amounts of C19 and C21 (Fig 4.8). Alkylresorcinols of etiolated leaf sheaths were also dominated by the C23 homolog, but together with C25 and C21. Etiolated stems had an alkylresorcinol distribution peaking at the C25 homolog, whereas the profiles of etiolated hypocotyls and leaf blades were relatively even across all major homologs including C25. Thus, the major difference between alkylresorcinol distributions on green and etiolated organs was the increased portion of the C25 homolog in the latter.   66                      Figure 4.8 Chain length distributions of normal alkylresorcinols in the waxes of various Secale cereale organs. (A) Green organs, (B) etiolated organs. The percentages of individual alkylresorcinol homologs are shown as mean values (n = 6) ± SD.   19 21 23 25 27010203040Green leaf bladeGreen external sheathGreen internal sheath19 21 23 25 27010203040Etiolated leaf bladeEtiolated sheathEtiolated stemEtiolated hypocotylRelative composition [% compound class]Alkylresorcinol chain lengthAB67  In B. distachyon, both the leaf blades and sheaths had alkylresorcinols dominated by the homolog with C19 alkyl chain (Fig 4.9), accompanied by the C21 homolog in approximately 2:1 ratios and minor amounts of C17, C23 and C25 homologs.             Figure 4.9 Alkylresorcinol chain length distributions in waxes on B. distachyon leaf blades and sheaths. The percentages of individual alkylresorcinol homologs are shown as mean values (n = 6) ± SD.   4.2.2.2 Methyl alkylresorcinol chain length distribution in Secale cereale and B. distachyon organs Individual methyl alkylresorcinols were quantified using GC-FID as described above for normal alkylresorcinols. In the Secale cereale etiolated leaf sheaths and etiolated stems, all methyl alkylresorcinols together accounted for ca 28% of the total AR mixture, in green external and internal sheaths and etiolated hypocotyls for ca 19%, and in green and etiolated leaf blades for ca 12% (Fig 4.10). Wax on green leaf blades contained mainly methyl alkylresorcinols with C19 (4.3%) and C21 (4.3%) alkyl chains, and methyl alkylresorcinol C21 Alkylresorcinol chain length17 19 21 23 25Relative composition [% compound class]0204060Leaf bladeSheath68  Methyl alkylresorcinol chain length19 21 23 250246810121416Green bladeGreen external sheathGreen internal sheathRelative composition [% compound class]19 21 23 250246810121416Etiolated bladeEtiolated sheathEtiolated stemEtiolated hypocotylABwas the predominant in wax from etiolated leaf blades (3.7%), green external (6.2%) and internal (7.6%) sheaths, etiolated sheaths (12.1%) and etiolated stems (8.8%). In contrast, wax from etiolated hypocotyls was dominated by the C19 homolog (7.9%). In B. distachyon, methyl alkylresorcinols were detected only in leaf blades, with alkyl chain lengths C19 and C21 in an approximate 2:1 ratio (Fig. 4.10).                   Figure 4.10 Chain length distributions of methyl alkylresorcinols in the total waxes on various S. cereale and B. distachyon organs. (A) Green S. cereale organs, (B) etiolated S. cereale organs, (C) leaf blades of B. distachyon. The percentages of individual methyl alkylresorcinol homologs are shown as mean values (n = 6) ± SD.   19 21024681012C69  4.2.2.3 Total cuticular alkylresorcinol amounts Alkylresorcinol quantities varied widely between waxes from different organs, and between etiolated and green tissues. Wax on green leaf blades of Secale cereale contained 0.34 ± 0.1 µg/cm2 of alkylresorcinols (Fig 4.11 A), corresponding to ca 2% of the wax mixture. Similarly, green external sheaths were covered with 0.38 ± 0.1 µg/cm2 (6% of total wax), etiolated stems with 0.39 ± 0.1 µg/cm2 (8% of total wax), and green internal sheaths with 0.33 ± 0.1 µg/cm2 of alkylresorcinols (6% of total wax). Relatively lower coverages were found for etiolated leaf blades 0.25 ± 0.1 µg/cm2 (2% of total wax), etiolated sheaths 0.26 ± 0.1 µg/cm2 (9% of total wax), and etiolated hypocotyls with less than 0.1 µg/cm2 (2% of total wax). Finally, in etiolated cotyledons no alkylresorcinols were detected. In B. distachyon, alkylresorcinols were detected only in leaf blades and sheaths (Fig 4.11 B), at levels of 0.8 ± 0.1 µg/cm2 (6% of the total wax) and 0.08 ± 0.01 µg/cm2 (1% of the total wax), respectively.            70             Figure 4.11 Total alkylresorcinol amounts. (A) Secale cereale organs in light and dark conditions, (B) B. distachyon organs. The coverages (µg/cm2) of alkylresorcinols are given as mean values (n=6) ± SD for the total wax extracted from entire organ.  Total alkylresorcinol amounts were determined by adding up quantities of all individual alkylresorcinol and methyl alkylresorcinol homologs detected by GC-FID.   4.2.3 Gene expression Two genes putatively involved in biosynthesis of alkylresorcinols, BdARS (from Brachypodium distachyon) and ScARS (from Secale cereale), had been previously isolated and partially characterized (Yao, 2011), including preliminary investigation of gene expression patterns. The present work aimed to provide further molecular evidence that these genes are indeed involved in formation of cuticular alkylresorcinols, by investigating the expression levels in all (etiolated or green) organs for which now also data on alkylresorcinol accumulation have been obtained. Semi-quantitative RT-PCR analyses showed that ScARS is expressed in all green organs (leaf blades, internal and external leaf sheaths) and in the Green bladeEtiolated bladeGreen ext sheathGreen int sheathEtiolated sheathEtiolated stemEtiolated hypocotylEtiolated cotyledon0.00.10.20.30.40.5Wax coverage [µ g/cm2 ]ABladeSheath C0.00.20.40.60.81.0B71  etiolated leaf blades and stems, but not in etiolated cotyledons of Secale cereale (Fig 4.12A). ScActin was used as a positive control, however it did not give a positive PCR product in samples from etiolated hypocotyls and leaf sheaths. BdARS was expressed only in leaf blades and sheaths of B. distachyon (Fig 4.12B). 18S rRNA positive control showed constitutive expression in all organs of this species.             Figure 4.12 Semi-quantitative RT-PCR analyses of gene expression. (A) ScARS in Secale cereale and (B) BdARS in B. distachyon. ScActin and 18S rRNA were used as constitutive controls in the analysis of ScARS and BdARS, respectively. Bl, blades; Hy, hypocotyl; Cy, cotyledon; Sh, sheath; exSh, external sheath; inSh, internal sheath; St, stem; iSt, internal stem; Sp, spike.   4.2.4 Subcellular localization. To also assess whether ScARS and BdARS have access to acyl CoA pools as putative substrates, the subcellular localization of the enzyme was investigated. To this end, ScARS and BdARS were fused with green fluorescent protein (GFP) either at the N- or C-terminus ScARS ScActin Bl Hy Cy Sh St Bl exSh inSh Etiolated Green Bl Sh St iSt Cy Sp BdARS 18S rRNA A B 72  for expression under the control of the constitutive cauliflower mosaic virus 35S promoter. Both constructs had previously been made by UBC Botany MSc student Ruonan Yao. The resulting constructs 35S:sGFP-ScARS or 35S:ScARS-sGFP and 35S:sGFP-BdARS or 35S:BdARS-sGFP were now transiently expressed in the abaxial epidermis of two week-old N. benthamiana leaves after agrobacterium-mediated infiltration. In addition, the ER-specific marker 35S:HDEL-RFP was co-infiltrated and expressed as an ER-specific marker.  Fluorescence signal was detected for both the N- and C-terminal fusions with GFP, indicating that protein expression was not silenced and that properly folded GFP was present. Since the C-terminal fusion gave stronger signal, 35S:ScARS-sGFP and 35S:BdARS-sGFP were chosen for detailed microscopy. Confocal laser scanning microscopy showed a reticulate pattern for the ScARS-GFP and BdARS-GFP fusion protein, indicating its localization in or near the ER (Fig 4.13). Merged images of ScARS and HDEL and BdARS and HDEL showed partial overlap between both signals, suggesting that both are not localized within the ER, but loosely associated with it.     73        Figure 4.13 Subcellular localization of ScARS and BdARS. (A) C-terminal fusion of ScARS with GFP (35S:ScARS-sGFP) (B) C-terminal fusion of BdARS with GFP (35S:BdARS-sGFP); (C) and (D) the ER-specific marker 35S:HDEL-GFP were transiently expressed in N. benthamiana epidermal cells; (E) and (F) merged image.    A  B C  D  E  F  74  4.3 Discussion The goals of this work were to analyze alkylresorcinols in the cuticular waxes on various organs of B. distachyon and Secale cereale grown in the light or dark, and to correlate the chemical data with expression patterns of two genes, BdARS and ScARS, putatively involved in formation of cuticular alkylresorcinols.   4.3.1 Identification of cuticular alkylresorcinols and methyl alkylresorcinols Alkylresorcinols with odd-numbered side chains ranging from C19 to C27 were first detected in S. cereale leaf blades, then also in waxes from green leaf blades, external and internal leaf sheaths, and from etiolated leaf blades, leaf sheaths, stems and hypocotyls. In addition, traces of even-numbered C20, C22, and C24 ARs were newly identified in all S. cereale organs except etiolated cotyledons, and homologs C29, C31 and C33 were detected only in etiolated hypocotyl. Moreover, a series of odd-numbered unsaturated ARs ranged between C19 to C27 were also detected in etiolated leaf blades and etiolated sheaths. Alkylresorcinols with odd-numbered side chains between C17 to C25 and C17 to C23 were found in B. distachyon leaf blades and sheaths, respectively. It is important to notice that ARs homolog detected in traces in this study have to be confirmed in further research.   The results presented here add even-numbered alkylresorcinol homologs to the chemical inventory of rye waxes, and they for the first time show the presence of similar series in various organs in both grass species. The new data further confirm the two previous reports on alkylresorcinols in plant waxes, where odd-numbered alkylresorcinols had been detected in the leaf blade waxes of Secale cereale and B. distachyon (Ji & Jetter, 2008; Yao, 2011). 75  Beyond these two wax studies, alkylresorcinols had been reported before in a large variety of plants (Kozubek & Tyman, 1999), but without distinguishing their specific location. Traces of even-numbered ARs had been reported only in barley and rye, but had again not been localized to the cuticular waxes (Briggs, 1974; Hengtrakul et al., 1991). Interestingly, the waxes of these two species had been analyzed repeatedly in earlier studies without identifying alkylresorcinols (Streibl et al., 1974; Tulloch & Hoffman, 1974).   In addition to the normal alkylresorcinols discussed above, four novel compounds with mass spectral characteristics similar to alkylresorcinols were found in the waxes of various Secale cereale organs. They were recognized as methyl derivatives of alkylresorcinols with C19, C21, C23 and C25 side chains based on two previous reports: similar methyl alkylresorcinol (MAR) homologs C19 to C27 had been reported once for wheat waxes, however without providing proof of structure or quantification (Adamski et al., 2013); methyl alkylresorcinols C19 and C21 had also been detected in B. distachyon leaf blade waxes before (Yao, 2011), again without giving details on the exact molecular structure.   Based on the characteristic fragment m/z 282 found in the mass spectra of all four putative MARs, it was concluded that they contained a methyl group on they benzyl carbon, in the 1’ position of the alkyl chain, or else on the aromatic ring in the ortho or para positions. To distinguish between isomers with side chain or ring methylation, the alkylresorcinol derivative bearing a methyl group in the position 1’ on a C21 alkyl side chain (CMAR) was synthesized. Its mass spectrum was identical with that of one MAR in rye wax, confirming the overall structure assignment of MARs (Fig 4.6). However, the GC elution behaviour 76  differed dramatically from that of the corresponding MAR in rye wax (Fig 4.7), ruling out that the wax constituent was side chain-methylated and instead suggesting an isomeric ring-methylated structure.  Unfortunately, attempts to also synthesize ring methyl ARs failed, so the exact position of the methyl group in the ring could not be determined. Further experiments are needed to distinguish between the ortho and para isomers, either using synthetic standards or isolating large enough quantities of MARs for NMR analysis.  Based on the chemical structure of MARs, the biosynthetic mechanisms leading to these products can be hypothesized. Overall, five alternative routes leading to MARs can be envisioned, falling into three categories. Firstly, 2-methyl-branched acyl CoA starter substrates might be incorporated, leading to CMARs (Fig. 4.14). This scenario can now be ruled out, since the wax MARs were found to have different isomeric structures. Secondly, straight-chain acyl CoAs might be elongated and cyclized into normal ARs, which then serve as substrates for SAM-dependent methyl transferases forming ring-MARs (Fig. 4.14). Precedence for this scenario comes from studies into Aspergillus terreus, where similar core MAR structures are formed by acetyl-CoA, malonyl-CoA and SAM-dependent (Matsuda et al., 2012). However, all previous reports are using Type I PKS, and it is not clear whether similar mechanisms may be operating in plants as well. Thirdly, straight-chain acyl CoA starters may be condensed with one methylmalonyl CoA and two malonyl CoA extenders, to form alkylresorcinols methylated in the ortho or para positions depending on extender incorporation sequence (Fig. 4.14). There is evidence that methylmalonyl CoA serves as extender in type III PKS reactions in plants and other organisms. For example, one of the chalcone synthase-like proteins identified in Pinus strobus showed a preference for 77  methylmalonyl CoA as extender u heterologous expression in E. coli (Schröder et al., 1998). Similarly, the bacterial SrsA from Streptomyces griseus (Funabashi et al., 2008; Nakano et al., 2012) and FtpA from Myxococcus xanthus (Hayashi et al., 2011) were found to biosynthesize ring MARs using methylmalonyl-CoA as extender in the first cycle of decarboxylative condensation reaction. In both species, the methyl groups were accordingly localized in the ortho position of the resorcinol ring.     78                 Figure 4.14 Methyl alkylresorcinol potential biosynthetic pathways.  The biosynthesis of MAR using (A) one molecule of malonyl-CoA and two molecules of malonyl-CoA as extender in different sequences with straight fatty acyl-CoA as starter substrate; (B) three molecules of malonyl-CoA as extenders and one molecule of methyl-branched fatty acyl-CoA as starter substrate; (C) ARs as substrate for SAM-dependent methyl transferases forming ring-MARs. H3C SCoAOnH3COn SCoAO O OH3COn SCoAO O OH3COn SCoAO O OH3COn SCoAO O OH3C n-1OHOHH3C n-1OHOHH3C n-1OHOHH3C n-1OHOHH3CCH3CH3CH3Methylmalonyl-CoAMalonyl-CoAMalonyl-CoAMethylmalonyl-CoA Malonyl-CoAMalonyl-CoAMethylmalonyl-CoAMalonyl-CoAMalonyl-CoAH3C SCoAOnCH3CH3CH3CH3CH3Malonyl-CoAMalonyl-CoA Malonyl-CoACO2CO2CO2CO2CO2CO2CO2CO2CO2CO2CO2 CO2CO2CO2CO2CO2H3C SCoAOnMalonyl-CoAMalonyl-CoA Malonyl-CoACO2 CO2CO2H3COn SCoAO O O CO2H3C n-1OHOHSAM-dependent methyl transferaseB A C 79  4.3.2 Homolog distributions of cuticular alkylresorcinols in Secale cereale and B. distachyon waxes The relative amounts of homologous alkylresorcinols differed between the wax mixtures from various organs of rye and B. distachyon, and also between green and etiolated tissues. On B. distachyon leaf blades, the alkylresorcinol homolog with C19 side chain dominated, followed by C21 and C17. This result matches a previous analysis of leaf wax from the same species (Yao, 2011). The present investigation adds to this knowledge by showing that alkylresorcinols in the wax from the B. distachyon leaf sheaths have a chain length distribution similar to that of leaf blades.  In Secale cereale, the chain length distribution of alkylresorcinols was also similar between all green organs, in this case showing a predominance of the C23 homolog, followed by C21 and C19. Again, these results confirm previous data on leaf blade alkylresorcinols (Ji & Jetter, 2008), and they add further information on other organs. In addition, alkylresorcinols even-numbered detected various organs, the specific and C29, C31 and C33 homologs found in etiolated hypocotyl, and odd-numbered unsaturated ARs homologs C19 to C27 detected in etiolated leaf blades and etiolated sheaths were not quantified in this study due to; firstly they were detected only in traces; secondly, some of them were detected in single samples only.   Interestingly, both grass species were thus found to contain alkylresorcinols with fairly similar chain length profiles on different organs. This result suggests that, on the one hand, the substrate pools available for alkylresorcinol biosynthesis have the same chain length composition in all organs of one species, and the acyl CoA-elongating enzymes must therefore be fairly similar throughout different parts of the plants. On the other hand, the results may also be explained by relatively strict substrate chain length specificities of the alkylresorcinol-forming enzymes, 80  suggesting that one and the same ARS is operating in all organs. The latter interpretation is supported by the relatively broad expression profiles of the candidate ARS genes also tested in this work (see 4.3.4). Conversely, the different alkylresorcinol chain length profiles in rye and B. distachyon waxes indicates that the ARS enzymes operating in both species may either have different chain length preferences, or else access and utilize acyl CoA substrate pools with characteristically different homolog compositions.  As a second major result, the chain length distributions of alkylresorcinols were found to differ greatly between green and etiolated organs of rye. Overall, the etiolated organs had a broader distribution of alkylresorcinol homologs, with lower amounts of the C23 homolog and increased levels of C21 and C25 homologs compared with corresponding green tissues. Based on the consistent expression of ScARS in green and etiolated tissues (see below), it seems plausible that the same alkylresorcinol-forming enzyme is operating independent of growth conditions. The differences in product chain length profiles then suggest that light conditions must affect the substrate pool available to the enzyme.   Overall, ARs chain length distribution had a similar effect than the cuticular waxes compound showed in the dark (Chapter 3), in which they had a broader distribution of homologs by compound class in etiolated organs. For instance in the dark, primary alcohol and free fatty acids had lower level of C26 homolog and increased levels of C22, C24 and C30; alkanes showed increased of C27 homolog and lower levels of C29 homolog, extending the spectrum of carbon chain length in each compound class (Fig 3.4 and 3.5). This difference in chain length 81  distribution suggests again that the biosynthesis of alkylresorcinol is part of wax biosynthesis machinery.  4.3.3 Total amounts of cuticular alkylresorcinols  The total amounts of cuticular alkylresorcinols were determined to correlate product amounts with gene activities in various organs of Secale cereale and B. distachyon. The waxes from green rye organs varied between 0.3 µg/cm2 and 0.4 µg/cm2, accounting for 2-6% of the total wax loads. These findings confirmed values reported previously for Secale cereale leaf wax (Ji & Jetter, 2008). In addition, the alkylresorcinol accumulation through the blades were demonstrated in the same study, where ARs are accumulated in a specific growth stage (15 com long, 12-14 days long), indicating also no presence of ARs at first two centimeters after POE, and then a peak at centimeters 4 to 10. Moreover, timing for wax production had been demonstrated that ARs were not formed at the same time as most other wax compounds (Ji, 2010), being biosynthesized later than other wax compounds.   On B. distachyon leaf blades, the total alkylresorcinol coverage was 0.8 µg/cm2, accounting for 6% of the total wax, again similar to a previous report (Yao, 2011). In contrast, leaf sheaths of this species had alkylresorcinol coverages a tenth of those on leaf blades. Contrary to Secale cereale, biosynthesis of alkylresorcinols in B. distachyon thus seems to be largely restricted to leaf blades.  Etiolation had varied effects on the amounts of cuticular alkylresorcinols on different rye organs. In particular, etiolated leaf sheaths and blades had similar alkylresorcinol amounts, and their coverages were reduced only slightly from corresponding green tissues. In contrast, etiolated 82  stems had relatively high alkylresorcinol concentrations, when compared with other etiolated organs, and etiolated hypocotyls and cotyledons had by far the lowest alkylresorcinol coverages. Unfortunately, the latter three organs could not be analyzed after growth in the light, due to technical limitations. All taken together, substantial amounts of alkylresorcinols were detected in all rye organs, independent of growth in the light or in the dark. This conclusion is contrary to previous studies, where rye and rice seedlings were found to contain higher concentrations of alkylresorcinols when grown in the dark (Deszcz & Kozubek, 2000;  Suzuki et al., 1996). However, those previous studies had quantified total plant alkylresorcinols, in units of µg/g, without normalizing for differences in plant size due to growth under low light stress and without discriminating between organs and tissues. In contrast, the current study focussed on cuticular alkylresorcinols and provided coverages measured in units of µg/cm2, accounting for differences in size and surface area of plants grown under various conditions.   4.3.4 Gene expression analysis and subcellular localization of ScARS and BdARS Two genes, potentially involved in the biosynthesis of cuticular alkylresorcinol, had been isolated from Secale cereale (ScARS) and B. distachyon (BdARS) and partially characterized (Yao, 2011). A former graduate student, Ruonan Yao, had also performed a partial gene expression study of these putative alkylresorcinol synthase genes, however that study was restricted to only a few plant organs due to time constraints. Here, semi-quantitative RT-PCR analysis was now employed to profile ARS gene expression in a wide range of organs, for comparison with the chemical results discussed above. ScActin was used a positive control for the gene expression analysis, however it did not give a positive PCR product for etiolated hypocotyls and leaf sheaths. Due to the underlying experimental problems with samples from 83  these two organs, ScARS expression cannot be assessed in them at this point and the gene expression tests will have to be repeated for these two organs. Of all other samples with positive controls in place, respective ARSs were expressed in those from green S. cereale organs, etiolated Secale cereale leaf blades, and B. distachyon leaf blades and sheaths. In contrast, gene expression was detected neither in etiolated Secale cereale cotyledons, nor in B. distachyon stems, cotyledons and spikes, coinciding with alkylresorcinol detection in wax analysis. Thus, the expression patterns of the candidate genes were found to, without exception, match the presence or absence of alkylresorcinol products in the various organs of both species and under light and dark conditions. These findings together provide important evidence supporting the hypothesis that ScARS and BdARS are involved in the biosynthesis of cuticular alkylresorcinols.  To further corroborate the involvement of the two ARS enzymes in the formation of cuticular alkylresorcinols, their subcellular localization was investigated. To this end, transient expression in Nicotiana benthamiana leaves via agrobacterium-mediated infiltration was employed, with HDEL as a counterstain specific for the ER membrane network. GFP fusion proteins of both ARS enzymes were found localized to a reticular ER-like network but not directly coinciding with the ER. Thus, the results indicate that both enzymes are loosely associated with the ER, which is the subcellular organ where VLC fatty acyl-CoA substrates are present and biosynthesis of cuticular wax is performed. This enzyme subcellular localization, together with previous evidence on the biochemical properties of the enzymes and the close correlation between ScARS and BdARS gene expression and organ-specific localization of cuticular alkylresorcinol, strongly suggests that ScARS and BdARS are indeed involved in the biosynthesis of cuticular alkylresorcinol. 84  Chapter 5: Alkyl ester compositions on green and etiolated organs of Secale cereale 5.1 Introduction Cuticular waxes consist of various very long chain alkanes, fatty acids, alcohols, ketones, aldehydes, and alkyl esters (Jetter et al., 2006). The wax alkyl ester amounts vary between plant species and organs, for instance on Copernicia cerifera leaves accumulating up to 85% wax ester (Kolattukudy, 1976). Simmondsia chinensis seeds had 97% of C38 to C44 wax esters (Benzioni & Forti, 1989).In contrast, Arabidopsis thaliana leaf wax contained only 0.1% to 0.2 % of alkyl esters, and stem wax ca 2.9% (Jenks et al., 1995). In Cosmos bipinnatus petals, alkyl esters were only detected in the adaxial cuticle (Buschhaus, 2010), whereas in jojoba (Simmondsia chinensis) they accumulated to very high concentrations on specifically seed storage lipid oils (Benzioni & Forti, 1989). Alkyl ester formation and accumulation can thus also be restricted to specific tissues only.  Beyond the total amounts of alkyl esters, also their homolog and isomer distributions are relevant to understand their biosynthesis and functions. Therefore, alkyl ester homolog distributions had been studied in many species such as Sorgum bicolor (Wettstein-Knowles & Mikkelsen, 1984) and Cyathea dealbata (Franich et al., 1985). However, the alkyl ester isomer composition has been studied only for a few plant species, originally employing wax fractionation, saponification and separate analyses of constituent acids and alcohols. These methods are time-consuming, and they yield only information on the overall homolog patterns of esterified acids and alcohols, but not within individual ester chain lengths. Therefore, a new method to identify and quantify alkyl ester isomers was proposed using gas chromatography-mass spectrometry (GC-MS) (Allebone & Hamilton, 1972) to assess acid from [RCO2H2]+ ions in the mass spectra. Using this approach, 85  extensive analyses of ester homologs in some species have been carried out (Sümmchen et al., 1995), providing valuable information on wax composition and biosynthesis. For instance, isomeric alkyl esters in Quercus robus contained a broad range of acid and alcohol moieties (Gülz et al., 1994), and more than 600 isomers were detected in wax esters from Cereus peruvianus (Dembitsky & Rezanka, 1996). Finally, the ester isomer compositions on inflorescence stem of Arabidopsis thaliana cer mutants has also been studied in detail (Lai et al., 2007), and all mutants and wild-type wax esters were found dominated by 16:0 acyl moieties.   Wax profiles, and alkyl ester homolog and isomer distributions in particular, can be used to infer biosynthetic mechanisms generating the complex mixtures with characteristic physical properties and associated biological functions. Wax biosynthesis starts with the elongation of saturated C16 and C18 fatty acyl CoAs to very-long-chain fatty acids (VLCFAs), thus generating chain length diversity ranging from 20 to 36 carbons. Subsequent modifications, either along the alcohol reduction pathway or the alkane decarbonylation pathway (Samuels et al., 2008) further generate compound class variety. Wax esters are formed by combination of acyl CoA elongation products and alcohols formed on the reduction pathway. Therefore, the overall ester chain lengths are  nearly double those of other wax compounds, typically ranging from 38 to 70 carbons (Jetter et al., 2006; Kunst & Samuels, 2003; Samuels et al., 2008).  Alkyl ester formation is catalyzed by three families of wax synthases (WSs) ( Jetter & Kunst, 2008); the first are mammalian WSs, first identified in mice, with high activity on C12-C16 acyl-CoAs and primary alcohols shorter than 20 carbons (Cheng & Russell, 2004). The second group are jojoba-type WSs, reported to use a large range of saturated and unsaturated C14-C24 acyl-86  CoAs, with highest activity on 20:1 acyl-CoA and 18:1 alcohol (Lardizabal et al., 2000). The third group of WSs, first identified in Acinetobacter calcoaceticus, is the enzyme family of WS/DGATs (Acyl-Coenzyme A:Diacylglycerol acyltransferases). They exhibit both WS and DGAT activity, thus can utilize both primary alcohols and more complex alcohols as substrates. WS/DGATs show a preference for C14 and C16 acyl-CoA substrates and C14 to C18 primary alcohols (Kalscheuer & Steinbüchel, 2003; Stöveken en al., 2005). In plants, ten WS/DGAT sequences have been annotated in Arabidopsis thaliana, and one of these (WSD1) has been characterized and determined to use C18, C24 and C28 alcohols and C16 fatty acyl-CoA to produce wax alkyl esters (Li et al., 2008). WSD1 is thus the only WS involved in the formation of cuticular wax esters that has been studied to date. However, since homolog and isomer distributions of wax esters vary greatly between plant species and organs, it is not clear whether similar WS/DGAT enzymes with widely differing substrate specificities may be responsible for generating diverse ester profiles, or whether WSs from other enzyme families are involved as well.  To enable further research into the biosynthesis of wax esters, homolog and isomer profiles of esters in the waxes of diverse species and organs have to be analyzed. Interesting targets for such investigations are the Poaceae, since waxes from various species in this group had previously been reported to contain high amounts of alkyl esters. For instance on Zea mays seedlings accumulating to 16% of the total wax (Bianchi et al., 1989) and on seven-day old Medicago sativa leaves to 11% (Avato et al., 1985). Previous research had also shown that Secale cereale waxes contained substantial amounts of esters (Ji & Jetter, 2008). However, rye ester compositions had been analyzed only qualitatively, and only for leaves grown in normal light 87  conditions (Ji & Jetter, 2008). The current project therefore aimed at a more comprehensive analysis of rye wax esters, in particular; i) to determine the chain lengths of alkyl esters in dark and light conditions, ii) to profile the ester isomers within each ester homolog, and iii) to compare the profiles of esterified alcohols and acids with those of free primary alcohols and fatty acids. To probe the variation of ester composition between organs and growth conditions, total wax was extracted and alkyl esters were analyzed from a variety of Secale cereale organs grown in the light (leaf blades, external and internal leaf sheaths) or dark (leaf blades, leaf sheaths, stems, hypocotyls and cotyledons). The alkyl ester isomer compositions in these waxes were determined and compared between selected green and etiolated Secale cereale organs.    88  5.2 Results The principal goal in this chapter was to analyze the composition of alkyl esters in waxes extracted from various Secale cereale organs grown in the light or dark. To this end, both the alkyl ester homolog distributions (5.2.1) and the isomer compositions within homologs were determined (5.2.2), and the results used to compare chain length distributions between total free and esterified alcohols and fatty acids (5.2.3).  5.2.1 Alkyl ester composition in waxes from Secale cereale organs Alkyl ester homologs were identified based on GC-MS characteristics using their retention order and molecular ions. To quantify the ester homologs, the corresponding peaks in GC-FID runs were compared with an internal standard, and total coverages of esters (5.2.1.1) and relative homolog amounts (5.2.1.2) calculated.  5.2.1.1 Total cuticular alkyl ester amounts To assess alkyl ester abundances in the Secale cereale waxes, homolog quantities were added together and compared with the amounts of other compound classes in respective wax mixtures (Table 5.1). Alkyl esters represented 10% (1.6 ± 0.4 µg/cm2) of the wax on green leaf blades, 12% (0.9 ± 0.1 µg/cm2) on green external sheaths and 12% (0.6 ± 0.1 µg/cm2) on green internal sheaths. Similarly, they accumulated to 6% (0.2 ± 0.1 µg/cm2) of the wax on etiolated sheaths, 6% (0.7 ± 0.2 µg/cm2) on etiolated leaf blades, 10% (0.5 ± 0.1 µg/cm2) on etiolated stems, 10% (0.3 ± 0.01 µg/cm2) on etiolated cotyledons and 17% (0.7 ± 0.2 µg/cm2) on etiolated hypocotyls.   89  Table 5.1 Compound class composition of waxes from various Secale cereale organs. Relative amounts [%] of compound classes are given as mean values (n = 6).    Secale cereale  relative composition [%]  Green organs  Etiolated organs  Leaf blades External sheaths Internal sheaths  Leaf blades Sheaths Stems Hypocotyls Cotyledons Fatty acids 1 2 6  1 2 7 16 11 Primary alcohol 66 36 43  61 32 36 17 9 Alkanes 3 7 5  1 2 2 3 2 Aldehydes 11 17 12  0 0 0 0 0 Alkyl esters 10 12 12  6 6 10 17 10 Alkylresorcinols 2 6 6  2 9 8 2 0 Not identified 7 20 16  30 49 38 46 68  5.2.1.2 Alkyl ester chain length distributions on green and etiolated Secale cereale organs Alkyl esters identified in waxes on all Secale cereale organs ranged from C38 to C52, with strong predominance of even-numbered homologs (Fig 5.1). The C46 alkyl ester homolog dominated in waxes on all green tissues, amounting to 35% of the ester fraction (0.6 ± 0.1 µg/cm2) on leaf blades, 37% (0.3 ± 0.03 µg/cm2) on external sheaths and 36% (0.2 ± 0.05 µg/cm2) on internal sheaths. Waxes from etiolated hypocotyls and cotyledons also had ester series dominated by the C46 homolog, with 40% and 42% of the fraction, respectively. Instead, the C44 homolog was prevalent on etiolated leaf blades (23% of the ester fraction), etiolated sheaths (19%) and etiolated stems (64%).  90  38es 40es 42es 44es 46es 48es 50es 52esWax coverage [µ g/cm2 ]0.00.20.40.60.8Green leaf bladesGreen external sheathsGreen internal sheathsEtiolated leaf bladesEtiolated sheathsEtiolated stemsEtiolated hypocotylsEtiolated cotyledons38         40        42         44         46         48         50        52 Figure 5.1 Chain length distributions of alkyl esters in the total waxes on various Secale cereale organs. The percentages of individual alkyl ester homologs are given as mean values (n=6) ± SD.  Waxes from all green organs had C52 alkyl ester as the second major homolog, comprising 14% of the fraction (0.2 ± 0.1 µg/cm2) on leaf blades, 20% (0.1 ± 0.01 µg/cm2) on external sheaths, and 19% (0.1 ± 0.02 µg/cm2) on internal sheaths. Waxes on etiolated leaf blades and stems had intermediate amounts of C52 ester (11% and 4% of the ester fractions, respectively), while other etiolated organs had relatively minor amounts of this homolog, comprising only 3% (≤0.1 µg/cm2) on sheaths, 2% on hypocotyls and <1% on cotyledons.   5.2.2 Chain length distributions of acyl and alkyl moieties in Secale cereale wax esters In order to analyze acyl and alkyl units within each of the wax ester homologs, GC-MS information was acquired. Chain lengths of esterified acids were assigned based on characteristic acyl ions [RCOOH2]+, with fragments m/z 229, 257, 285, 313, 341, 369, 397, 425 and 453 91  indicating the presence of even-numbered C14 to C30 acids, respectively. Respective ion counts were used to calculate relative abundances of all acyl moieties, and the amounts of accompanying alkyl moieties were assigned accordingly. Finally, the overall amounts of esterified acid and alcohol homologs were calculated by multiplying individual ester homolog coverages with isomer percentages.  The wax mixtures on various Secale cereale organs contained alkyl esters composed of primary alcohols ranging from C16 to C34, with the C26 homolog predominant throughout (Fig 5.2). Wax esters on green and etiolated leaf blades had 72% (0.7 ± 0.1 µg/cm2) and 58% (0.2 ± 0.06 µg/cm2) of C26 alcohol, respectively. Additionally, C22 and C24 alcohols were found esterified in relatively high abundances, on green leaf blades at 6% and 12%, and on etiolated leaf blades at 20% and 17%, respectively. Taken together, these data show that etiolated leaf blades had wax esters with lower percentages of C26 alcohol and increased amounts of C22 and C24 alcohols compared with green leaves. The same trend was also observed for leaf sheaths, with 61% (0.4 ± 0.1 µg/cm2) of C26 esterified alcohol on external green sheaths and only 27% (≤ 0.1 µg/cm2) on etiolated leaf sheaths, again accompanied by corresponding changes in C22/C24 alcohol percentages.  92  Esterified alcohol chain length16 18 20 22 24 26 28 30 32Relative composition [% ]020406080Green leaf bladesGreen external sheathsGreen internal sheathsEtiolated leaf bladesEtiolated sheaths Figure 5.2 Chain length distributions of esterified alcohols in the waxes from various Secale cereale organs. Waxes from green leaf blades, external and internal sheaths, etiolated leaf blades and sheaths were analyzed. Percentages of homologs are given as mean values (n=6) ± SD.  The chain length distributions of esterified acids differed drastically from the patterns found for esterified alcohols. The dominant esterified fatty acid homolog differed between ester homologs (Table 5.2), and variation in ester chain lengths was thus largely due to variation in chain length of the acids bound to C26 alcohol. As an exception to this overall trend, all (green and etiolated) organs had C38 alkyl ester composed of relatively high amounts of C16 fatty acid and C22 alcohol. Similarly, all organs except for green leaf blade also had C40 alkyl ester containing C16 fatty acid esterified with C24 alcohol. Etiolated sheaths had a distinctive ester isomer composition, not driven primarily by combination of C26 alcohol with various acids. Instead, ester homologs up to C42 were found to contain mainly C14 to C20 acids, while C44 and higher esters contained mainly C22 to C28 fatty acids. Accordingly, corresponding ester alcohols varied between C20 and C28.    93  Table 5.2 Relative amounts [%] of acyl moieties in wax esters from various Secale cereale organs.    Chain length C38 ester C40 ester C42 ester C44 ester C46 ester C48 ester C50 ester C52 ester Green leaf blades 14 18.5 ± 9.8 52.8 ± 8.2 0.4 ± 0.1 0.2 ± 0.01 0.3 ± 0.01 0.2 ± 0.1       16 30.1 ± 14.9 23.9 ± 6.0 86.1 ± 4.3 2.9 ± 0.6 0.5 ± 0.1 0.4 ± 0.1 0.2 ± 0.3 0.1 ± 0.2 18 5.9 ± 3.3 4.6 ± 1.3 6.2 ± 1.7 70.3 ± 5.1 1.4 ± 0.4 0.8 ± 0.2 0.2 ± 0.3 0.2 ± 0.3 20 28.1 ± 14.0 16.2 ± 5.2 6.3 ± 2.5 23.7 ± 5.2 95.1 ± 0.4 9.2 ± 1.3 2.3 ± 1.0 0.6 ± 0.5 22 0.7 ± 0.8 2.4 ± 1.6 0.6 ± 0.2 1.9 ± 0.5 1.5 ± 0.3 85.0 ± 2.3 5.3 ± 0.9 1.2 ± 0.7 24       0.1 ± 0.2 0.2 ± 0.0 0.4 ± 0.0 3.0 ± 1.2 77.2 ± 5.0 2.5 ± 0.7 26          0.3 ± 0.1 0.1 ± 0.0 0.7 ± 0.3 14.4 ± 4.5 94.0 ± 1.4 28          0.3 ± 0.1 0.7 ± 0.1 0.8 ± 0.2 0.3 ± 0.4 1.4 ± 0.7  0.01 ± 0.00 0.03 ± 0.01 0.20 ± 0.07 0.22 ± 0.06 0.56 ± 0.15 0.22 ± 0.06 0.12 ± 0.04 0.22 ± 0.07 Green external  sheaths 14 5.2 ± 0.8 13.2 ± 4.6 0.8 ± 0.1 0.3 ± 0.01 0.3 ± 0.01 0.2 ± 0.1 0.2 ± 0.2 0.2 ± 0.1 16 82.7 ± 1.8 64.6 ± 3.3 78.8 ± 3.8 10.8 ± 1.7 0.7 ± 0.1 0.9 ± 0.3 0.7 ± 0.4 0.3 ± 0.2 18 7.1 ± 2.0 11.3 ± 0.7 7.5 ± 1.4 60.7 ± 4.7 1.4 ± 0.3 0.9 ± 0.3 1.1 ± 0.5 1.0 ± 1.1 20 2.1 ± 1.2 8.3 ± 0.8 10.7 ± 2.2 23.3 ± 2.5 93.4 ± 1.1 7.9 ± 1.4 1.6 ± 0.5 1.0 ± 0.7 22 0.5 ± 0.4 0.8 ± 0.4 0.8 ± 0.1 2.5 ± 0.5 1.4 ± 0.4 78.5 ± 3.4 4.9 ± 1.9 1.0 ± 1.2 24 1.2 ± 1.0 0.8 ± 0.3 0.4 ± 0.2 0.8 ± 0.1 0.9 ± 0.2 5.3 ± 0.8 71.6 ± 2.3 1.3 ± 0.8 26 0.2 ± 0.2 0.7 ± 0.1 0.4 ± 0.1 0.8 ± 0.3 0.6 ± 0.2 4.0 ± 0.8 15.5 ± 1.4 91.2 ± 3.3 28 0.2 ± 0.2 0.1 ± 0.1 0.6 ± 0.01 0.7 ± 0.1 1.2 ± 0.01 2.1 ± 0.2 4.0 ± 0.8 3.8 ± 1.2  0.00 ± 0.00 0.02 ± 0.01 0.09 ± 0.03 0.10 ± 0.02 0.31 ± 0.04 0.10 ± 0.03 0.07 ± 0.02 0.17 ± 0.01 Green internal sheaths 14 5.4 ± 2.2 7.3 ± 1.4 0.6 ± 0.1 0.4 ± 0.1 0.3 ± 0.0 0.3 ± 0.1 0.5 ± 0.1 0.2 ± 0.1 16 83.3 ± 2.3 72.1 ± 4.0 73.0 ± 8.3 9.7 ± 1.7 1.0 ± 0.3 0.9 ± 0.2 0.8 ± 0.4 0.4 ± 0.1 18 5.4 ± 1.8 10.3 ± 1.6 7.8 ± 0.9 52.9 ± 12.0 1.4 ± 0.2 0.7 ± 0.2 0.9 ± 0.3 0.4 ± 0.1 20 2.1 ± 0.3 6.8 ± 1.8 16.8 ± 7.0 30.7 ± 8.7 92.2 ± 2.1 7.2 ± 1.3 1.0 ± 0.4 0.6 ± 0.1 22 1.5 ± 1.2 2.0 ± 2.7 0.5 ± 0.1 4.1 ± 1.6 2.7 ± 1.2 75.8 ± 3.5 4.0 ± 2.7 0.6 ± 0.6 24 0.9 ± 0.8 0.6 ± 0.1 0.5 ± 0.1 1.0 ± 0.3 0.8 ± 0.6 9.5 ± 2.8 70.3 ± 1.8 2.1 ± 0.9 26 0.2 ± 0.3 0.6 ± 0.2 0.3 ± 0.0 0.6 ± 0.1 0.6 ± 0.2 3.6 ± 1.0 19.5 ± 2.2 92.0 ± 0.8 28 0.6 ± 0.5 0.1 ± 0.1 0.5 ± 0.0 0.5 ± 0.0 1.1 ± 0.1 1.7 ± 0.3 2.9 ± 0.4 3.5 ± 0.3  0.01 ± 0.00 0.02 ± 0.00 0.06 ± 0.02 0.07 ± 0.03 0.22 ± 0.05 0.07 ± 0.02 0.05 ± 0.01 0.11 ± 0.02 Etiolated leaf blades 14 3.5 ± 2.6 14.9 ± 6.3 0.2 ± 0.4 0.1 ± 0.2 0.1 ± 0.1 0.2 ± 0.3 0.2 ± 0.3 0.1 ± 0.2 16 85.4 ± 3.8 39.7 ± 2.9 78.7 ± 2.8 2.2 ± 0.8 0.2 ± 0.3 0.2 ± 0.4 0.5 ± 0.7 0.1 ± 0.2 18 9.6 ± 3.1 34.8 ± 5.6 10.8 ± 1.2 69.2 ± 5.2 1.5 ± 0.7 0.3 ± 0.3 0.4 ± 0.7 0.4 ± 0.6 20 0.5 ± 0.9 9.6 ± 0.9 8.9 ± 1.9 23.5 ± 3.8 89.1 ± 3.0 2.4 ± 2.6 0.7 ± 1.0 0.2 ± 0.3 22 0.3 ± 0.4 0.3 ± 0.3 1.1 ± 0.8 3.9 ± 1.4 3.8 ± 0.9 63.5 ± 7.4 2.1 ± 2.1 1.5 ± 1.2 24 0.4 ± 0.6 0.4 ± 0.6 0.1 ± 0.1 0.6 ± 0.5 3.6 ± 0.7 9.1 ± 2.1 50.7 ± 5.9 0.3 ± 0.6 26 0.1 ± 0.2 0.2 ± 0.3 0.2 ± 0.3 0.3 ± 0.3 1.4 ± 1.1 23.5 ± 5.4 41.7 ± 3.1 96.1 ± 3.3 28 0.1 ± 0.2 0.1 ± 0.1 0.1 ± 0.2 0.1 ± 0.2 0.3 ± 0.3 0.7 ± 0.9 3.5 ± 4.4 0.9 ± 1.4  0.03 ± 0.01 0.06 ± 0.02 0.14 ± 0.04 0.15 ± 0.05 0.11 ± 0.02 0.06 ± 0.02 0.03 ± 0.01 0.08 ± 0.02 Etiolated sheaths 14 10.2 ± 1.3 8.6 ± 3.8 1.2 ± 1.1 0.8 ± 0.7 1.8 ± 2.0 1.4 ± 0.6 3.4 ± 2.3 0.5 ± 0.2 16 74.2 ± 3.9 42.9 ± 3.5 34.5 ± 0.9 8.2 ± 1.8 5.0 ± 3.1 5.3 ± 3.1 4.0 ± 1.5 0.2 ± 0.3 18 9.3 ± 1.7 32.6 ± 6.2 9.8 ± 1.7 14.9 ± 1.7 5.9 ± 1.2 2.5 ± 0.4 3.7 ± 1.5 1.4 ± 0.3 20 1.2 ± 0.2 12.3 ± 3.8 22.2 ± 3.3 17.1 ± 3.6 20.6 ± 2.7 5.4 ± 2.7 6.7 ± 3.1 4.8 ± 1.5 22 0.8 ± 1.4 2.1 ± 1.4 14.4 ± 3.7 20.4 ± 2.1 19.0 ± 2.5 26.2 ± 3.3 6.8 ± 2.6 1.6 ± 0.4 24 1.2 ± 2.0 1.0 ± 1.1 16.3 ± 3.1 37.0 ± 1.8 36.9 ± 4.7 33.0 ± 6.0 43.2 ± 4.2 8.5 ± 4.1 26 1.1 ± 1.0 0.2 ± 0.3 0.7 ± 0.6 0.8 ± 0.3 10.1 ± 2.1 22.9 ± 3.1 22.4 ± 4.9 69.8 ± 8.2 28 1.7 ± 1.9 0.0 ± 0.0 0.4 ± 0.4 0.4 ± 0.4 0.5 ± 0.6 2.5 ± 2.1 9.5 ± 5.5 10.5 ± 5.1   0.01 ± 0.00 0.02 ± 0.00 0.03 ± 0.00 0.03 ± 0.00 0.03 ± 0.00 0.02 ± 0.00 0.02 ± 0.01 0.01 ± 0.00                            Predominant isomers are highlighted in bold    94  5.3 Discussion The objective of this chapter was to identify and quantify alky esters in the cuticular waxes on various organs of Secale cereale grown in the light or dark, including detailed analyses of homolog and isomer distributions within the ester mixtures.  5.3.1 Total alkyl ester amounts in waxes from Secale cereale organs Alkyl esters were detected as a major compound class in the cuticular waxes from all Secale cereale organs. Green leaf blades had 10% of alkyl esters in the total wax, matching values of 10-11% reported previously for Secale cereale leaves (Ji & Jetter, 2008; A. P. Tulloch & Hoffman, 1974). Green sheath wax had a slightly higher percentage of alkyl esters compared with green blades, while etiolated blades and sheaths had relatively low amounts instead. These data taken together suggest that etiolation of rye leaves resulted in reduced levels of alkyl esters, in contrast to a previous report on increased ester levels in etiolated barley leaves (Giese, 1975; Macey, 1970).  Alkyl esters had been previously detected in a great variety of species (Allebone & Hamilton, 1972; Griffiths et al., 2000; Gülz et al., 1994; Jetter & Riederer, 1996; Lai et al., 2007; Shepherd et al., 1995), at varying levels in different organs. For instance, Simmondsia chinensis seeds had 97% of C38 to C44 wax esters (Benzioni & Forti, 1989), leaf wax of Copernicia cerifera contained 85% of esters (Kolattukudy, 1976), and Arabidopsis thaliana leaf wax only 0.1% of esters (Jenks et al., 1995).   95  5.3.2 Alkyl ester chain length distribution in green and etiolated Secale cereale organs The homologous series of alkyl esters ranged between C38 to C52 in waxes from Secale cereale organs grown in the light or dark. Previously, only alkyl ester chain lengths C40 to C48 had been reported for Secale cereale leaf wax, and only qualitative information on the homolog distribution had been provided (Ji & Jetter, 2008). The C46 homolog was predominant in all green organs selected, confirming prior information on Secale cereale leaves (Tulloch & Hoffman, 1974). However this result is in contrast to another study by Ji and Jetter (2008), where C44 had been reported as the predominant homolog in Secale cereale leaf wax. Additionally, green organs had C52 alkyl ester as a second major homolog. Thus, the ester homolog distribution is varying drastically between species as, for instance, Picea abies esters contained 30% of the C40 homolog (Sümmchen et al., 1995), Papaver rhoeas and Eschscholzi californica had C44, Papaver bracteatum, Papaver dubium and Papaver orintale had C46, and Papaver somniferum had C48 as the predominant homolog (Jetter & Riederer, 1996).  It had been reported that overall alkyl ester amounts may differ between green and etiolated organs (Giese, 1975; Macey, 1970), however there was no information on the homolog distributions in the dark and light. In the present study, C44 ester dominated on etiolated leaf blades, etiolated sheaths and etiolated stems, and C46 on etiolated hypocotyls and cotyledons. These data, in comparison with chain length distributions on green organs, suggest that etiolation prompted a shift toward shorter ester chain lengths.   96  5.3.3 Acyl and alkyl moieties in wax esters from selected green and etiolated Secale cereale organs Beyond of the analysis of alkyl ester total amounts and chain length distributions, also the isomer composition of esterified alcohol and acid moieties on Secale cereale organs was investigated here for first time. Only leaf blades and sheaths were selected for this analysis, due to limited reproducibility of the material harvested from other organs. The isomer analysis showed that C26 esterified alcohol was predominant in all Secale cereale organs. In other species, esterified alcohol homologs with other chain lengths had been prevalent, for instance C22 alcohol in Rudus idaeus flowers and C24 in Crataegus monogyna flowers (Griffiths et al., 2000). However, esterified hexacosanol had been reported as predominant on green and etiolated H. vulgare leaves (Giese, 1975), and on Lolium perenne and Chenopodium album leaves (Allebone & Hamilton, 1972).   Esterified fatty acid chain lengths varied between ester homologs, thus complementing C26 esterified alcohol to form various overall ester chain lengths. A similar trend had been reported for Hordeum vulgare (Giese, 1975) and Lolium perenne (Allebone & Hamilton, 1972), where alcohols were also dictating alkyl ester structures. In contrast, esters on Vicia faba (Griffiths et al., 1999), R. idaeus and Crataegus monogyna flowers (Griffiths et al., 2000), and A. thaliana stems (Lai et al., 2007) had been found to contain one predominant esterified fatty acid together with various alcohols, suggesting that in these species ester formation was fairly specific for acid substrate chain length.   97  5.3.4 Chain length distribution of free and esterified alcohol and fatty acids To further assess substrate specificities of ester formation in rye, the chain length distributions of esterified (this chapter) and free alcohols (see chapter 3) can be compared between green and etiolated organs. It was found that amounts of C26 esterified alcohol decreased in etiolated leaf sheaths and blades, whereas C22 and C24 esterified alcohols increased in comparison with green organs (Fig 5.3).                 Figure 5.3 Chain length distributions of free and esterified alcohols in the cuticular waxes on various Secale cereale organs in light and dark conditions. Percentages of homologs are shown as means (n=6) ± SD.  Green blade16 18 20 22 24 26 28 30 32020406080100Etiolated leaf blades16 18 20 22 24 26 28 30 32020406080100Green external sheaths16 18 20 22 24 26 28 30 32020406080100Green internal sheathsCarbon chain length16 18 20 22 24 26 28 30 32020406080100Etiolated sheaths16 18 20 22 24 26 28 30 32020406080100Free alcoholsEsterified alcoholsGreen leaf bladesRelative composition [%]98  Most interestingly, the esterified alcohol patterns in each organ and light regime showed similar patterns as the accompanying free alcohols, suggesting that the pool of free alcohols, formed on the reduction pathway, serves as substrates for wax alkyl ester biosynthesis.  The same conclusion had also been drawn from studies on Hordeum vulgare leaves (Giese, 1975) and Arabidopsis thaliana stems (Lai et al., 2007).  The chain length distributions of esterified and free fatty acids differed between waxes from various organs. Esterified fatty acids tended to have shorter chain lengths than free fatty acids in the same organs. Thus, for example the fatty acid up to C26 constituted ca 98% of the ester acyls, and only very low amounts of > C26 chain lengths were esterified in all organs. In contrast, free fatty acids on all organs had chain length distributions comprising larger amounts of C26 and higher homologs (chapter 3). Furthermore, etiolated organs were found to have ester acyl distributions fairly similar to corresponding green organs. Overall, these results suggest that ester formation occurs with a certain acid specificity, with preference for acyl CoA substrates containing 26 or fewer carbons and discriminating against longer chains.          99  Green blade14 16 18 20 22 24 26 28 30 32 340102030405060 Etiolated leaf blades14 16 18 20 22 24 26 28 30 32 340102030405060 Green external sheaths14 16 18 20 22 24 26 28 30 32 340102030405060 Green internal sheathsCarbon chain length14 16 18 20 22 24 26 28 30 32 340102030405060 Etiolated sheaths14 16 18 20 22 24 26 28 30 32 34Relative composition [%] 0102030405060Free fatty acidsEsterified fatty acidsGreen leaf blades            Figure 5.4 Chain length distributions of free and esterified fatty acids in the cuticular wax of various Secale cereale organs in light and dark conditions. Percentages of homologs are shown as means (n=6) ± SD.             100  Chapter 6: Conclusions and future directions Based on previous studies, it was hypothesized that secondary metabolites such as alkylresorcinols are targeted to the cuticle for specific ecophysiological functions, that incorporation of very-long-chain aliphatic structures may render them hydrophobic and thus miscible with cuticular waxes, and that such aliphatics may be derived from fatty acid biosynthesis using VLC acyl-CoAs formed during wax production. As a prerequisite for future studies into the ecophysiological functions of alkylresorcinols, the present work investigated their chemical structures and biosynthesis in two model grass species. Some major conclusions can be drawn from comparisons of wax compositions on different organs grown either in the light or in the dark, in particular on the mechanisms underlying wax formation (6.1), on the enzymes responsible for alkylresorcinol biosynthesis (6.2), and on novel methyl alkylresorcinols (6.3).   6.1 The wax biosynthesis on various organs of Secale cereale and Brachypodium distachyon This work has provided detailed information on the wax compositions of various organs of the two model grass species S. cereale and B. distachyon that now enable detailed comparisons.  For example, it was found that in both species leaf blades had drastically higher wax coverages than sheaths. These findings suggest that wax deposition occurred mainly at or near the point of emergence (POE) between sheath and blade, thus confirming previous notions. The cuticular waxes on all organs had similar compound class compositions dominated by primary alcohols, despite variations in total wax amounts between organs. Moreover, aldehydes and primary alcohols shared similar homolog distributions peaking at C26, leading to the conclusion that both compound classes are biosynthetically related. Wax aldehydes thus serve as intermediates en 101  route to primary alcohols, contrary to Arabidopsis thaliana, where aldehydes occur as intermediates in the biosynthesis of wax alkanes instead. Finally, the isomer analysis of alkyl esters showed that also esterified alcohols were dominated by the C26 homolog in all S. cereale organs, indicating that the same pool of free alcohols may serve as substrate for alkyl ester formation or for export to the cuticle (Fig 6.1)    Further comparisons between organs grown in the light or in the dark showed that etiolation caused reduced total wax amounts and a shift to longer and/or shorter chain lengths, hence overall broader homolog distributions. Interestingly, aldehydes were not detected on etiolated organs, whereas alkanes were found at relatively high concentrations in these waxes. This finding further confirms that these two compound classes are formed independently in the investigated grass species, different from the commonly accepted model for alkane formation via aldehyde intermediates in A. thaliana.            102                      Figure 6.1 Potential Cuticular wax biosynthetic pathways on grasses. The core represents C16 fatty acyl-CoA undergo successive elongations, extending the acyl chain by C2 units. The VLCFAs wax precursors are then used for the production of aldehydes, primary alcohols and alkyl esters (left), and alkylresorcinols and alkanes (right). C26 primary alcohol is the central key compounds for alkyl esters formation. The biosynthesis of aldehydes is proposed as intermediate in the biosynthesis of primary alcohols. Cuticular alkylresorcinols are biosynthesized using VLCFAs as substrate.  C16 CoAC263 x malonyl CoA CO2ARSC52C46C40 C24C22C18 CoAC20 CoAC22 CoAC24 CoAC26 CoAC28 CoAC30 CoAC32 CoAC263 x malonyl CoA CO2ARS3 x malonyl CoA CO2ARS3 x malonyl CoA CO2ARS3 x malonyl CoA CO2ARS3 x malonyl CoA CO2ARSC22C28C30C18C20C24C28C30C32C50C48C44C42C38C17C19C21C23C25C27C29C27C25C26C28C30C32C24103  6.2 Biosynthesis of cuticular alkylresorcinols Alkylresorcinols were detected on all S. cereale organs selected (except on cotyledons), demonstrating a broad distribution of ARs throughout all areal surfaces of this species. In contrast, ARs were detected only in two organs of B. distachyon, suggesting a much more specific profile of AR biosynthesis gene activity in this species. The expression patterns of the previously isolated candidate ARS genes closely matched the presence or absence of alkylresorcinol products in the various organs of both species and under light and dark conditions.  In addition, subcellular localization of the corresponding proteins indicate that both enzymes are loosely associated with the ER, which is the subcellular organ where VLC fatty acyl-CoA substrates are present and biosynthesis of cuticular wax is performed. These findings together provide important evidence supporting the hypothesis that ScARS and BdARS are involved in the biosynthesis of cuticular alkylresorcinols. In addition, based on the expression of ScARS in green and etiolated tissues, it seems plausible that this ARS operates independent of growth conditions. Consequently, shifts in AR chain length profiles between green and etiolated organs must be due to changes in substrate pools available for ARs biosynthesis. On the other hand, similar effects of etiolation were also found for other wax compounds on the various organs, indicating that both VLC aliphatics and alkylresorcinols shared the same acyl-CoA intermediates. Altogether, the current findings therefore support the hypothesis that cuticular alkylresorcinols are derived from epidermal fatty acid biosynthesis, formed on a branch pathway of wax biosynthesis.   104  So far, there is no evidence for the occurrence of ARs in A. thaliana, neither from our lab nor from the vast literature on this model species. Thus, it can be assumed that A. thaliana does not biosynthesize ARs, and offers a clean background for biochemical tests using heterologous expression of genes involved in AR formation. Vectors harboring the ScARS and BdARS genes can transformed into A. thaliana using Agrobacterium and floral dip techniques. Assuming that the corresponding enzymes gain access to the wax precursor pools and that the AR products are exported from the ER to the cuticule, this system will then allow investigations into the specificity of the ARSs. In particular, A. thaliana mutants, such as cer6 and cer4, with varying compositions of VLC acyl-CoA pools as substrate may offer the opportunity to test the substrate preference profile of these two genes. First, it has been demonstrated that CER6 encodes an elongase condensing enzyme required for the synthesis of VLCFAs precursors (Fiebig et al., 2000; Millar et al., 1999). Accordingly, the cer6 mutant accumulated C24 and C26 CoAs (Jenks et al., 1995). Second, the CER4 gene encodes a fatty acyl-CoA reductase involved in alcohol-forming pathway. Thereby, cer4 mutant has drastically reduced levels of primary alcohols and wax esters and increased aldehyde levels (Jenks et al., 1995), suggesting an increase in the flux of fatty acyl precursors into the decarbonylation pathway. Consequently, this mutant likely has increased acyl CoA amounts available for parallel pathways such as AR formation, making it another good candidate for testing ARS functions and specificities. In the present research work, stable transformation of both Arabidopsis mutants was attempted, however the results were not satisfactory and therefore not presented in this thesis. Instead, the experiment has to be repeated in order to test substrate preference profile of the two ARS genes by expression in the Arabidopsis cer6 and cer4 mutants.  105  6.3 Methyl alkylresorcinols in grasses A new group of alkylresorcinol derivatives were detected for first time in various organs of S. cereale, and identified as methyl alkylresorcinols. Three potential isomers were hypothesized according to their mass spectral characteristics. Analysis of a synthetic standard ruled out that the methyl group is localized at the benzylic position, suggestion that the methylation should instead be in the resorcinol ring. However, it can currently not be determined whether the methyl group is attached in the para or ortho position. Hypothetical biosynthesis mechanisms are proposed involving either methylmalonyl-CoA as extender or SAM-dependent methylation. All these possibilities have to be tested using synthetic standards with specific methylations in the ring, and/or feeding experiments with labeled methylmalonyl-CoA as extender. Further experiments must test whether ScARS and BdARS are involved in the biosynthesis of MARs, or whether other ARSs may show the necessary activity towards methylmalonyl-CoA extenders. In particular, crystallization of the enzyme(s) and X-ray crystallographic determination of its (their) three-dimensional structure could explain the substrate specificity of the ARSs, and gene-silencing techniques might be employed to suppress ARS expression and test whether cuticular ARs and/or MARs are affected.   In conclusion, the current study provided detailed chemical information on the wax amounts and compositions on various organs of S. cereale and B. distachyon grown in the light or the dark. Specifically, alkylresorcinol homolog distributions were investigated, to contribute information towards a better understanding of the biosynthesis of waxes in grass species and the biosynthesis of cuticular ARs. In addition, MARs were reported for the first time in S. cereale, opening opportunities for further research into the mechanism of ARS enzymes. Finally, the 106  characterization of BdARS and ScARS reveled that they are likely responsible for the biosynthesis of cuticular alkylresorcinols, thus providing new tools to investigate the biological function of ARs in grasses in the future.                     107  References Adamski, N. M., Bush, M. S., Simmonds, J., Turner, A. S., Mugford, S. G., Jones, A., … Uauy, C. (2013). 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