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Alkylresorcinols in the leaf cuticular wax of secale cereale l Ji, Xiufeng 2010

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ALKYLRESORCINOLS IN THE LEAF CUTICULAR WAX OF SECALE CEREALE L. by  XIUFENG JI B.Sc. and M.Sc. Central China Normal University, 2002  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE STUDIES (Chemistry)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) April 2010  Xiufeng Ji, 2010  Abstract Alkylresorcinols (ARs) are bioactive compounds found in 11 plant families. Indirect evidence showed that they were likely near/at the surface of plant organs, suggesting their defensive role against biotic and abiotic stressors in the environment. However, neither the exact function nor the forming mechanisms are known. To assess whether their primary function is exerted at plant surface and to unravel their biosynthesis, the local distribution of ARs has to be determined first. Hence, the goals of the research were to analyze the ARs in the leaf waxes, to compare their amounts with interior concentrations, to determine AR distribution within the surface wax layers, and to monitor their accumulation as a function of leaf development. Rye (Secale cereale L.) was chosen for this investigation, since previous studies had indicated that it has high levels of ARs in various organs. The total wax mixture firstly extracted consisted of primary alcohols (71%), alkyl esters (11%), aldehydes (5%), and small amounts of alkanes, steroids, secondary alcohols, fatty acids and unknown compounds. ARs were identified by GC-MS and comparison with nonadecylresorcinol (AR19:0). They contributed 3% of total wax, and comprised a homologous series with odd-numbered alkyl side chains from C19 to C27. Secondly, abaxial and adaxial waxes separately sampled, contained very similar relative quantities of all constituents. Thirdly, the epicuticular and intracuticular wax layers were selectively extracted. ARs comprised 2% of the intracuticular wax, yet none in the epicuticular wax. All other wax components were spread uniformly between both wax layers. By analyzing various segments at four growth stages of rye leaves, the spatial distribution of waxes along the length of leaf blade and wax accumulation over time were assessed. All the major compound classes shared similar wax production zone, spatial distribution pattern and timing for wax production. Yet ARs were formed in a remarkably different spatial area and time periods. They were only detectable at later growth stages (IV and III) and not detected near POE in contrast to major wax constituents which were produced from each growth stage to its next adjacent stage and mainly within 2-cm segments beyond POE.  ii  Table of Contents Abstract………………...……………………………………………………………………….ii Table of Contents…………………...………………………………………………………… iii List of Tables…………………………………………...………………….…………………...vi List of Figures…………………………...……………………………….……………………vii Abbreviations………………………….………………………………….……………………ix Acknowledgements……………………………...……………………….……………………..x Co-authorship Statement…………………………..………..………………………………..xi  Chapter 1: Introduction and literature review……………..…….………..……………………..1 1.1 Plant cuticular waxes.………………………………………….…………………………...1 1.1.1 Wax organization…………………………………………...…..….………..................1 1.1.2 Wax function……………………………………………..…………..…….…..............1 1.1.3 Wax composition…………………………………....…………..………….….............2 1.1.4 Wax biosynthesis………………………………………………..………...…...............3 1.2 Plant alkylresorcinols.…………………………………..…........……..………………….5 1.2.1 Occurrence of alkylresorcinols in plants…..………………………………..................5 1.2.2 Bioactivities of alkylresorcinols……………………………………..………...............6 1.3 Cuticular alkylresorcinols.…………………………………………..…..…………………7 1.3.1 Occurrence of alkylresorcinols in the plant cuticle……………………..…..................7 1.3.2 Alkylresorcinol bioactivity in the plant cuticle………………………………...............7 1.4 Biosynthesis of alkylresorcinols in plants.……… ………………..………………………9 1.4.1 Biosynthetic pathway to alkylresorcinols…………………………..……….................9 1.4.2 Enzymes or genes discovered responsible for alkylresorcinol biosynthesis…………12 1.5 Thesis objectives.…………………………………..…………………………………...….13 References.……………………………………………………………………………………..14  Chapter 2: Alkylresorcinol deposition in the cuticle along rye (Secale cereale) leaves ……………………………………….……………….…………...…………………….17 2.1 Introduction………………………………………………………..………………………17 2.2 Materials and methods……………………………………………………………………17  iii  2.2.1 Plant materials and growth conditions……………..………………………................17 2.2.2 Wax extraction………………………………………………….…………….............18 2.2.3 Synthesis of 5-n-tridecylresorcinol (6, AR13:0) ……………………………………18 2.2.4 Synthesis of 5-n-nonadecylresorcinol (1, AR19:0) ………………………………….19 2.2.5 Sampling ARs from internal tissue of rye leaves……...……….………….................20 2.2.6 Chemical analysis…………………………………………………….………............20 2.3 Results and discussion…………………………………………………….…….………...22 2.3.1 Total wax……………………………………………………………..….……...........22 2.3.2 Identification of alkylresorcinols in cuticular wax………………….………..............26 2.3.3 Alkylresorcinols in internal tissue …………………………………..……………….28 2.3.4 Abaxial and adaxial wax…………………………………………………...…............28 2.3.5 Epi- and intracuticular wax……………………………………………...……............29 2.4 Conclusion…………………….….………………………………………………………..33 References……………………………..………………………………………………….…....34  Chapter 3: Time course of alkylresorcinol deposition in the cuticle along the rye (Secale cereale) leaf………………………………………………………………………………….…36 3.1 Introduction ……………………………………………………..……………………..….36 3.2 Materials and methods………………...………………………………………………….37 3.2.1 Plant materials and growth conditions……………..……………………………........37 3.2.2 Wax extraction……………………………………………….…………………….....39 3.2.3 Chemical analysis…………………………………………………….………….....39 3.3 Results and discussion……………..……………………………………….………….….40 3.3.1 Leaf growth over time………………………………………………….……………..40 3.3.2 Total waxes…………………………………………………………………………...42 3.3.2.1 Spatial distribution of total waxes……………………………...………………42 3.3.2.2 Accumulation of total waxes over time………………………….……………..43 3.3.3 Compound classes…………………………………………………………………….44 3.3.4 Chain length distribution………………………………………………..……………48 3.4 Conclusions………………………….……………………………………………………..50 References……………………………..………………………………………………….……51  iv  Chapter 4: Conclusions and future work ………………………………..………...……………52 References……………………………..………………………………………………….…....55 Appendix A: Supporting information for time course………………….…………..…......……56 Appendix B: Cloning of alkylresorcinol synthase (ARS) genes……….…………..……………65 B.1: Summary of cloning procedures………………………………...………………………65 B.1.1 Strategies for ARS gene cloning………………………………………………..……65 B.1.2 Procedures of ARS gene cloning……………………………………………….……67 B.2: Problems encountered……………………………………………………………...……68 Reference………………………………………………………………………………………69  v  List of Tables Table 2.1 Very long chain compounds identified in rye leaf waxes…….…………………….……24 Table 2.2 Homolog and isomer composition of very long chain alkyl esters identified in rye leaf waxes….…………………………………..……………………………………………………24 Table A.1 Statistical comparison of the accumulation of total wax at four growth stages. Listed data are P values at N=6 by one-way ANOVA……….…………………………………….………60 Table A.2 Statistical comparison of the accumulation of primary alcohols at four growth stages. Listed data are P values at N=6 by one-way ANOVA…………………………………………61 Table A.3 Statistical comparison of the accumulation of secondary alcohols at four growth stages. Listed data are P values at N=6 by one-way ANOVA…………..……………………..………62 Table A.4 Statistical comparison of the accumulation of alkyl esters at four growth stages. Listed data are P values at N=6 by one-way ANOVA…..…………………………………….………63 Table A.5 Statistical comparison of the average wax loads of alkanes at four growth stages. Listed data are P values at N=6 by one-way ANOVA………….……………………………..………64 Table B.1 Primers designed based on the consensus of STS/CHS from peanut (Arachis hypogaea), grape (Vitis vinifera) and Arabidopsis (Arabidopsis thaliana)…………………..……..……65 Table B.2 Primer designed based on the consensus of STS/CHS from rice (Oryza sativa)…….….65 Table B.3 Primer designed based on the consensus of three ARS from rice (Oryza sativa) and two from sorghum (Sorghum bicolor)………………………….………………………………...…66 Table B.4 Primer designed based on the consensus of blasted sequences using the two sorghum (Sorghum bicolor) ARS sequences to rye sequence database..……………………………...…66  vi  List of Figures Fig. 1.1 Cross section showing view of plant cuticle……...………………………………….…1 Fig. 1.2 Biosynthetic pathways to typical wax components. (A) Elongation of the fatty acyl chain by four enzymes in one cycle. (B) Overall biosynthetic pathways…………………..3 Fig. 1.3 Structure of alkylresorcinols (ARs)…………………………………………………..…5 Fig. 1.4 Isotope labeling experiments..…………………….………………………………...…10 Fig. 1.5 Proposed mechanism for AR biosynthesis…...………………………………..………11 Fig. 2.1 Synthesis of 1,3-dihydroxy-5-nonadecylbezene (AR19:0)……………………………19 Fig. 2.2 Derivatization reaction with N,O-bis(trimethylsilyl)-trifluoroacetamide……..………21 Fig. 2.3 Compound class composition of cuticular wax mixtures from the second leaf of rye…...………………………………………………………………………………….…23 Fig. 2.4 Chain length distributions of individual wax compounds in the total, adaxial and abaxial wax mixtures on the second leaf of rye…..…………………….…………………25 Fig. 2.5 Structures of alkylresorcinols identified in rye leaf wax (1-5) and synthesized as standards for structure elucidation (1) as well as quantification (6)…..…….……..…...…26 Fig. 2.6 Chain length distributions of alkylresorcinols in the total, adaxial and abaxial wax mixtures on the second leaf of rye…….………………………………………….….……27 Fig. 2.7 Relative amounts of wax compound classes within the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye……….…………………………..…30 Fig. 2.8 Chain length distributions of individual wax components in the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye…..…………...…31 Fig. 2.9 Chain length distributions of alkylresorcinols in the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye…..……………………………….…32 Fig. 3.1 Schematic overview of the sampling design employed in this study….………………38 Fig. 3.2 Growth of the second leaf of rye….………………………………………………...…41 Fig. 3.3 Distribution of total wax amounts along the second leaf of rye at four growth stages…...……………………………………………………………………………….…42 Fig. 3.4 Distribution of individual wax compound classes along the second leaf of rye at four growth stages…..……………………………………………………………………….…45 Fig. 3.5 Relative amounts of compound classes in rye leaf waxes….…………………………46 Fig. 3.6 Chain length distributions within compound classes in rye leaf waxes…..………...…48  vii  Fig. 3.7 Percentage of alkylresorcinol homologs within the alkylresorcinol fractions..…….…49 Fig. A.1 Percentages of compound classes in total wax at various growth stages……………..56 Fig. A.2 Percentage of primary alcohol homologs in total primary alcohols…………...……...57 Fig. A.3 Percentage of individual alkane homologs and fatty acid homologs in total alkanes and total fatty acids, respectively…………………………………………..………………….58 Fig. A.4 Percentages of individual ester homologs in total esters…………………..………….59  viii  Abbreviations 9-BBN  9-borabicyclo[3.3.1]nonane  ANOVA  analysis of variance  AR  Alkylresorcinol  ARS  Alkylresorcinol synthase  BSTFA  N,O-bis(trimethylsilyl)-trifluoroacetamide  CC  column chromatography  cDNA  complementary deoxyribonucleic acid  CHS  chalcone synthase  CoA  coenzyme A  DNA  deoxyribonucleic acid  dNTP  deoxynucleotide triphosphate  EST  Expressed Sequence Tag  FA  fatty acid  FID  flame ionization detector  Fig.  figure  g  gram  GC  gass chromatography  mL  microliter  m/m  mass/mass  mRNA  messenger ribonucleic acid  MS  mass spectrometry  PCR  polymerase chain reaction  PKS  polyketide synthase  POE  point of emergence  SD  standard deviation  STS  Stilbene synthase  THF  tetrahydrofuran  TLC  thin layer chromatography  TMSi  trimethylsilyl  USDA  United States Department of Agriculture  UV  ultraviolet  VLCFA  very long chain fatty acid  WS  wax synthase ix  Acknowledgements I would like to express my deep and sincere gratitude to my supervisor Dr. Reinhard Jetter, the guardian of principles, whose understanding, personal guidance, patience and tolerance have not only provided a good basis for the present thesis, but also helped me through personal growth. Thank you to past and present members of the Jetter lab for helpful discussions and advice. Especially Stephen Greer, Christopher Buschhaus, Miao Wen, Clare van Maarseveen, Zhonghua Wang, and Bangjun Wang, I could not have been here without you guys. Thank you to Stillman, Lisa, Vera Burnham, Qian Wang, Suzanne and Bob Taylor for all the help and love along the way. Thank you to the UBC Chemistry department for funding. Lastly, and most importantly, thank you to my husband, Zhan, for his love, tolerance and support.  x  Co-authorship Statement This project was initiated by my supervisor, Prof. Dr. Reinhard Jetter. Through the literature searching and preliminary investigation, I selected the model plant for the study. In Chapter 2, I conducted all the extractions and chemical analysis, and prepared the first version of the manuscript. Dr. Jetter assisted in revision and finalized the manuscript for publication. For Chapter 3, I did all the experiments and collected all the data. Dr. Jetter helped in designing the styles of presenting data. Under his supervision, I wrote the first version of the manuscript and revised it many times before he started to work on the final version for publication. This work has produced one published paper and the other one to be published. Through all stages of the project, Dr. Jetter has provided great suggestions and counseling.  xi  Chapter 1: Introduction and literature review  1.1 Plant cuticular waxes 1.1.1 Wax organization The surface of primary above-ground plant organs is covered by a cuticle, consisting of cutin and cuticular waxes. Cutin is the major component of the cuticle (40-80% of cuticle mass). It is a three-dimensional polyester lattice formed by interlinked hydroxy fatty acids (usually 1618 carbons long). This cutin network is both impregnated and covered with cuticular waxes. The embedded cuticular wax is intracuticular wax. The thin, continuous wax film covering the surface of cutin is epicuticular wax, which forms the outermost layer of cuticle (Fig.1.1).  Fig. 1.1 Cross section showing view of plant cuticle. * Diagram is not to scale (modified from Schaffer et al. 2000).  1.1.2 Wax function Plant cuticular wax exists as the interface between plant tissue and the environment, and therefore plays pivotal physiological and ecological roles in a plant’s life. It is generally accepted that cuticular wax is the major barrier to prevent non-stomatal water loss across the plant surface.1 Experiments on tomato fruit surface without stomata have shown that the aliphatic constituents in the intracuticular wax layer form the main portion of the water-loss barrier, whereas epicuticular aliphatics play a minor role.2 Plant surfaces covered with wax crystals have self-cleaning property (often called Lotus-effect), hence protecting the surface 1  against dirt adhesion. Wax crystals increase the hydrophobic surface area, making the plant surface highly water repellent and therefore the contact area between the water droplet and plant surface is very small. Hence adhesion between the droplet and particle occurs when a droplet rolls over particles, in which way the particle is removed and the plant surfaces are protected against accumulation of dust and air pollutants.3,4 Barthlott and Neinhuis’ studies on artificial surfaces with different roughness showed that surface roughness is essential for hydrophobic surfaces to execute the self-cleaning function.3 In addition to those two physiological roles, the cuticle plays a key role in protecting the plant from ultraviolet radiation.5,6 Holmes and Keiller found that epicuticular wax crystals scatter part of the radiation to protect plants against ultraviolet radiation.7 Ecologically, epicuticular wax has been shown to affect plant-insect interactions, e.g., the attachment of insect feet to plant surfaces.8-11 Other studies have shown that reduction in cuticular waxes can increase susceptibility to pathogens thereby suggesting that cuticular wax is also involved in plant defense against bacterial and fungal pathogens.8,12  1.1.3 Wax composition Cuticular waxes are complex mixtures of very long chain fatty acid (VLCFA) derivatives, alicyclic and aromatic components. VLCFA derivatives are most often unbranched and fully saturated alcohols, aldehydes, ketones, fatty acids, esters and alkanes. Each of these compound classes comprises a homologous series with chain lengths ranging from 20 to 36 carbons, except esters from 36 to 70.13 Waxes also include cyclic compounds such as triterpenoids and phenylpropanoids.14 The composition and quantity of cuticular wax can vary widely not only among plant species, but also among different organs of a single species, and even for the same organ at different growth stages. Wax analysis of needles of Taxus baccata revealed a gradient of nonacosan-10-ol from the inner parts of the cuticle to the outer layers, and showed that cyclic and relatively polar components tend to accumulate in intracuticular wax.15  2  1.1.4 Wax biosynthesis Wax biosynthesis starts with the elongation of fatty acyl chains to form VLCFAs. The elongation is catalyzed by fatty acid elongation enzyme complexes. Starting with the preexisting C16 or C18 acyl-CoA, one two-carbon unit is added in each elongation cycle comprising four reactions (Fig.1.2A): (1) condensation of malonyl-CoA with a long-chain acylCoA to form a β-keto acyl-CoA; (2) reduction to a β-hydroxyacyl-CoA; (3) dehydration of the hydroxyl group to an enoyl-CoA; (4) reduction of the enoyl-CoA to elongated acyl-CoA. Only even-carbon-number fatty acid intermediates are produced after each elongation cycle. Among the four enzymes involved in one elongation cycle, 21 condensing enzyme-like gene sequences have been identified in the Arabidopsis thaliana genome; β-keto acyl-CoA reductases in Zea mays (maize) and in Arabidopsis thaliana, an enoyl reductase gene in Arabidopsis thaliana, and a very-long-chain hydroxy fatty acyl-CoA dehydratase in Arabidopsis thaliana have been identified and characterized.16-19 (A)  (B)  O R  C18 CoA  SCoA O HO  O  Fatty Acid Elongation  SCoA  (1)Condensing enzyme  C26  CO2 O  C28  O  R  SCoA  Acyl Reduction Pathway  C30  (2)β-keto acyl reductase  C32 O  OH O  De novo Elongation  R R  Decarbonylation Pathway  SCoA  n  O OH  R  (3)Dehydratase  O R  O R  OH  n  n  H  SCoA R  (4)Enoyl reductase  n  OH  R n  Monooxygenase O R  SCoA  Esters  Secondary alcohols + Ketones  Fig.1.2 Biosynthetic pathways to typical wax components. (A) Elongation of the fatty acyl chain by four enzymes in one cycle. (B) Overall biosynthetic pathways.  3  VLCFAs formed by elongation cycles are used for biosynthesis of wax components. For most plants, there are two major wax biosynthetic pathways (Fig.1.2B): the acyl reduction pathway yielding even-numbered (C20 to C36) fatty acids, primary alcohols and esters (C36 to C70), and the decarbonylation pathway, producing odd-numbered (C19 to C35) alkanes, secondary alcohols and ketones.13 Alcohol-generating reductase in pea leaves and in Arabidopsis thaliana have been purified and characterized.20,21 The final step of the acyl reduction pathway is the synthesis of wax esters catalyzed by a wax synthase (WS). One WS gene has been cloned from jojoba embryo and another WS in Arabidopsis thaliana was identified and characterized.22,23 The decarbonylation pathway is initiated by reduction of VLCFAs to aldehydes. Odd-numbered alkanes are thought to be formed after elimination of carbon monoxide in the presence of aldehyde decarbonylase. Vioque and Kolattukudy have purified and characterized an aldehyde-generating reductase in pea leaves.20  4  1.2 Plant alkylresorcinols 1.2.1 Occurence of alkylresorcinols in plants Alkylresorcinols (ARs), also called 1,3-dihydroxy-5-alkylbenzenes, 5-alkylresorcinols, or resorcinolic lipids (Fig.1.3), have been found in plants, fungi, animals and bacteria. OH 2 3  1 4  6 5  HO  R  Fig. 1.3 Structure of alkylresorcinol (AR).  In plants, ARs occur as homologous series with alkyl chain lengths ranging from five to 29 carbons.24 In most cases, the side chain in ARs is odd-numbered, which is significant with regard to their biosynthetic pathway. For each of these homologues, the alkyl chain of ARs may be saturated, monounsaturated, or polyunsaturated. The side chains may also be modified by keto and hydroxy groups. Besides chain-modified derivatives, in some cases, ARs have ringmodified (2,4-alkyl) derivatives or hydroxy-modified derivatives (such as, hydroxy groups replaced by methoxy goups). ARs have been found in eleven plant families: Anacardiaceae, Ginkgoaceae, Proteaceae, Myrsinaceae,  Primulaceae,  Myristicaceae,  Iridaceae,  Compositae,  Leguminosae  and  Gramineae.24 ARs were initially isolated and characterized in fruits, seeds, and all senescent organs, and later in green organs such as leaves and stems. The amounts of ARs in plants vary largely depending on the source. The highest concentrations of ARs have been reported for the oil extracted from the cashew nut shell, which contains up to 20% (m/m) of ARs.24 Other plant sources contain amounts of ARs that vary from 0.005% (m/m) for Hordeum vulgare (barley) grains to 0.3% (m/m) for Secale cereale (rye) grains.25-27  5  1.2.2 Bioactivities of alkylresorcinols Due to the presence of the separate hydrophilic (dihydroxybenzene ring) and lipophilic (long aliphatic chain) regions in AR molecules, ARs have amphiphilic properties and therefore have effects on biological membranes. For example, it has been shown that ARs, especially the unsaturated homologs, induce an increased permeability of the bilayers of liposomes for ions and small non-electrolytes.28 Other reported biological activities of ARs include effects on the activity of nucleic acids, enzymes and cells, and growth regulation. For instance, ARs extracted from the leaves of Oncostemon bojerianum are known to exhibit cytoxic activity against the A2780 ovarian cancer cell29 and another study has found the main cause of the decreased growth of animals fed with AR-containing cereal grains is related to an AR-induced decrease of the appetite, though the mechanism of this process is not yet known.24 Although ARs are found in increasing numbers of organisms, a broader understanding of their bioactivities and the underlying mechanisms is lacking.  6  1.3 Cuticular alkylresorcinols 1.3.1 Occurrence of alkylresorcinols in the plant cuticle Indirect evidence shows that ARs accumulated at or near the surface of various organs. Ross et al. have demonstrated that almost all ARs are present in the bran (composed of cell layers and all exposed at the surface of cereal grains) and shorts (a mixture of bran and flour) fractions of wheat and rye grains, whereas white wheat flour has essentially no ARs present and rye flour has only a low amount.25,26 Extracting slices of fruit peel (1-2 mm thick) of Mangifera indica (mango) in 95% ethanol, Droby and Prusky reported AR fractions of 190 µg/g fresh weight in fruit peel, equivalent to twelve times of that which is in the flesh (15 µg/g fresh weight). More interestingly, 96 h after peeling, the outer layer of mango flesh accumulated AR fractions at concentrations of 160 µg/g fresh weight while the flesh below did not.30 ARs have also been found only in the seed covers of Myristica fragrans (nutmeg, ethanol extracts) or shell of Anacardium occidentale (cashew nut).24,31 Since grain bran, seed covers, nut shells, fruit peels, and outer layers of peeled fruits are all plant surfaces, it can logically be inferred that ARs occur specifically at plant surfaces. Furthermore, since plant surfaces are the place where cuticular waxes reside, it is quite likely that ARs specifically appear in the plant cuticle. To date there is one report on the occurrence of ARs in the cuticular wax of barley (Hordeum vulgare) seeds extracted by dipping seeds in CHCl3 for 30 seconds,32 but unfortunately the cuticular waxes of diverse other species known to synthesize ARs have not been investigated for AR occurrence. Thus, direct evidence for surface accumulation of ARs is scarce.  1.3.2 Alkylresorcinol bioactivity in the plant cuticle ARs exposed at the plant surface have been found to have antibacterial and antifungal activity, and cause contact dermatitis. AR fractions in the cuticular waxes of barley seeds are responsible for resistance against pathogenic fungi such as Aspergillus niger and recent experiments have shown antibacterial effects of ARs extracted from seed covers of Myristica fragrans (nutmeg).24,32 It has also been reported that the unpeeled mango (Mangifera indica) fruits are resistant to Alternaria alternate, but freshly peeled fruits are susceptible because of the antifungal compounds, 5-(12-cis-heptadecenyl)-resorcinol and 5-pentadecylresorcinol 7  which are at fungitoxic concentrations in the peel, but not in the flesh of unripe mango fruit. In this same study, the flesh of peeled mango fruits also became resistant to Alternaria alternate infections within 24 h after peeling due to accumulation of ARs in the outer layer of the flesh.30 Diogenes et al. have described 5-pentadecylresorcinol-induced dermatitis among cashew nut workers who have direct skin contact with raw nut shells, while the dermatitis did not or less happen to workers dealing with peeled nuts or heated unpeeled nuts (heat is to diminish the time spent in removing the shells).32 Reffstrup et al. have shown that Philodendron-induced dermatitis is caused by the presence of 5-n-heptadecenylresorcinol.33 All of these findings suggest that ARs may occur specially at plant surfaces where cuticular waxes are deposited but we still lack definitive knowledge as to whether ARs have biological functions based solely on their bioactivities in the plant cuticles, in contrast to that which they may have in internal tissues. The existence of ARs near the surface of some plant species and organs, plus their lipophilic properties of alkyl side chains and their antibacterial functions, brings up the question whether the ARs are located in the cuticular waxes of those systems. Cuticular ARs have been described only once in seeds of barley (Hordeum vulgare) cultivars,32 with no detailed homolog profiles mentioned. Furthermore, no other AR-containing species have been analyzed for whether ARs exist in cuticular waxes. For those AR-synthesizing plants, their wax mixtures were not reported to contain ARs. For instance, rye (Secale cereale L.) grains and seedlings were reported to include relatively high amounts of ARs,34,35 but ARs were not reported in the cuticular waxes of leaves and straws from this species.36,37 It might be due to the organs (leaves and straws) which do not contain ARs yet other AR-containing organs (grains and seedlings) were not analyzed. Therefore, it is not clear whether ARs in those examples accumulated partly or entirely in the cuticular wax layer, and whether they are aimed at the cuticle to play biological functions at the plant surface. It is also not apparent where ARs might be located throughout epi- and intracuticular layers.  8  1.4 Biosynthesis of alkylresorcinols in plants 1.4.1 Alkylresorcinol Biosynthesis Isotope labeling experiments using labeled malonate or acetate have established that the AR ring is formed by an elongation plus cyclization biosynthetic pathway (Fig.1.4A, 4B).38,39 For instance, Dayan et al. fed either [2-13C] acetate or [2-13C] malonate separately to the seeds of Sorghum bicolor × Sorghum sudanense hybrid, and found in  13  C NMR spectra a 2,4,6  isotope pattern (Fig.1.4A) in the ring of sorgoleone extracted from seedlings, indicating that AR15:3, the precursor of sorgoleone, was formed via a tetraketide intermediate through a polyketide pathway (Fig.1.4A).  38  Besides, [1-13C]-acetate was found to be incorporated into  positions C1 and C3 of the aromatic moiety of sorgoleone in the root exudates of Sorghum bicolor, confirming the polyketide nature of the AR ring.39 Based on these findings, it has been suggested that AR formation combines elongation and cyclization as shown in Fig.1.5. In the proposed scheme, condensation of an acyl-CoA with three malonyl-CoAs is thought to be catalyzed by a polyketide synthase to yield a linear tetraketide intermediate, without reductive removal of oxygen as in fatty acid biosynthesis. Then an aldol-type cyclization followed by decarboxylation results in the AR ring. The reaction is similar to that catalyzed by stilbene synthases (STS), rather than Claisen condensation catalyzed by chalcone synthases. The variety of polyketides found in nature arises from the incorporation of different starters during biosynthesis. For example, 6-methylsalicylic acid is formed with acetyl-CoA as a starter (Fig.1.4D). Chalcone synthesis depends on p-coumaroyl-CoA. Analogously, it is expected that the formation of ARs with different chain lengths depends to a high degree on the specificity of the appropriate enzyme for the starters. The 6-methylsalicylic acid synthetase isolated from Penicillium patulum showed much higher specificity for acetyl-CoA than propionyl-CoA (which acted as the starter at 13% of the rate of 6-methylsalicylic synthesis in the presence of acetyl-CoA) and almost no activity with butyryl-CoA or hexanoyl-CoA.40 However, a polyketide synthase extracted from Cannabis sativa was suggested to be responsible for olivetol (AR5:0) biosynthesis from hexanoyl-CoA and malonyl-CoA.41  9  (A) O  O  Acetyl-CoA ligase  Acetyl-CoA carboxylase  CoA-S  HO  Acetate  O  O  CoA-S  Acetyl-CoA  OH  Malonyl-CoA  CoA-S  O  3×  O R  O  Acyl-CoA  -CO2 -H2O R  O  O  OH HO  S-Enzyme  Tetraketide intermediate  O  OH  H3C-O O  (B)  OH  O  Sorgoleone  O OH  HO  HO  H3C-O O O O  (C) O CoA-S  HO  SCoA  2×  R  O  CO2  O  Reduction Dehydration O  O  R S-Enzyme  O R S-Enzyme  Malonyl-CoA  1× CO2  R=CH3, C11H23, C13H27, C15H31, C16H33, C17H35, C17H33 and C17H31  -H2O O  O  O  HO  R  S-Enzyme  R COOH  6-Alkylsalicylic acids  (D)  CoA-S  O  O O  O  HO R  SCoA  3×  O  O  O  OH  -H2O -CO2 -HSACP  CO2 R  SACP  HO  R  5-Alkylresorcinols  R=C9H19-C19H39, C15H29 and C17H33  Fig. 1.4 Isotope labeling experiments. (A) and (B) were designed to explore the ring formation in AR biosynthesis while (C) and (D) were to find out the starters. In (A), [2-13C] acetate and [2-13C] malonate were used respectively. In (B), [1-13C] acetate was used. (C) used methyl esters of [1-14C] fatty acids or [1-14C] acetate and (D) used [1-13C] fatty acyl esters. (A) and (B) showed that AR ring is formed through a polyketide pathway while (C) and (D) suggested the fatty acyl- or acetyl-CoAs to be the starters. Compared with AR biosynthesis, anacardic acid  10  biosynthesis involves the partial reduction and dehydration, and no decarboxylation after aldoltype cyclization. Note: Acyl carriers such as CoA, ACP and Enzyme are written based on the proposed pathways in the papers cited.  O O R  HO  SCoA  O SCoA  O  CO2  3× 3¡Á  R CoAS  OH  O  O O  - CO2 - HSCoA - H 2O  R  OH  Fig. 1.5 Proposed mechanism for AR biosynthesis. Suzuki et al. fed rice (Oryza sativa) seeds with 1-13C labeled fatty acid esters and found the pre-existing fatty acyl-CoAs incorporated into AR molecules (side chain, 9≤C≤19) in seedlings (Fig.1.4D).42 Among the fatty acid starters (FA12:0, FA13:0, FA15:0, FA17:0 and FA19:0) fed at the same concentration, the enzyme in rice plants tends to transform preferentially FA13:0 to the corresponding AR12:0. Labeling experiments using 1-14C fatty acids (FA12:0, FA14:0, FA16:0, FA17:0, FA18:0, FA18:1, FA18:2 and FA18:3) methyl esters showed the incorporation of them into anacardic acids in inflorescences of Pelargonium xhortorum (Geranium) (Fig.1.4C). One cultivar of the Geranium incorporated mostly FA18:0 into corresponding anacardic acids, while the other converted similar amounts of FA18:0, FA18:2 (ω 6, 9) and FA17:0 into anacardic acids.43 For ARs with side chains longer than C19, no labeling experiments have been tried yet, perhaps due to the very low solubility of very long chain fatty acids and their esters.  11  1.4.2 Enzymes or genes discovered responsible for alkylresorcinol biosynthesis As mentioned before, STSs share the same mechanisms of elongation and cyclization with the enzyme proposed to be involved in AR biosynthesis, though STSs use p-coumaroylCoA as the natural starter, instead of fatty acyl-CoAs for AR biosynthesis. Thus, the two enzymes are expected to have the same catalytic sites, similar residues and topology along active site pockets. It is believed that alkylresorcinol synthases (ARS) belong to type III polyketide synthase (PKS) family, like stilbene synthases (STSs). Type III PKS enzymes are structurally and mechanistically distinct from the type I (modular type) and type II (subunit type) PKSs. Type II and type III PKSs are from plants and bacteria while type I PKSs from yeast and animals. Type III PKSs use acyl-CoAs as substrates to carry out a series of condensation, cyclization, aromatization and decarboxylation reactions within a single active site.  Although the overall mechanism for AR biosynthesis has been proposed, only a few ARSs involved in AR biosynthesis have been characterized. The first alkylresorcinol synthase, ArsB and its gene were reported responsible for n-heneicosylresorcinol (AR21:0) in the bacterium Azotobacter vinelandii by Funa and co-workers. ArsB accepted acyl-CoAs from C10 to C22, yet showed highest activites for C18, C20 and C22 acyl-CoAs. Another alkylresorcinol synthase, srsA from bacterium Streptomyces griseus was found to use branched C15 and C16 acyl-CoAs as starter substrates and condenses extender units in the strict order of malonyl-CoA, malonyl-CoA, and methylmalonyl-CoA, resulting in methyl-substituted alkylresorcinols. Besides the two ARSs found in bacteria, five ARSs were discovered in plants by researchers at USDA, within which two were from sorghum and three from rice (Baerson, pers. communication). Yet ARS genes responsible for the biosynthesis of cuticular alkylresorcinols have not been reported to date.  12  1.5 Thesis objectives The appearance of ARs on the surface of some plant species, together with their lipophilic properties and antibacterial functions, raises the questions whether ARs are located in those cuticular waxes, whether ARs accumulate partially or completely in the cuticular wax layer, and whether they are intended for biological functions at the plant surface. The first two questions can be addressed by chemical analysis. As for the last one, the localization of the enzymes synthesizing ARs may give hints of whether ARs are meant for cuticular wax. Cloning the gene via mRNA is prerequisite, which means the information about when mRNA is expressed is necessary. The timing can be best assessed by a time course of cuticular AR accumulation. Thus the objectives of the project were set to address the questions raised. Work was focused on the following investigations: (1) To examine the chemical composition of leaf cuticular wax and find out whether ARs exist in the cuticle. (2) To find out to what extent ARs accumulate in cuticle in contrast to inner tissues. (3) To investigate the chemical composition of epi- and intracuticular waxes and locate ARs in cuticular wax. (4) To monitor cuticular wax over the course of leaf development and find out when and where ARs are deposited on the plant surface Rye (Secale cereale L) was chosen as the model plant for the study after preliminary investigations had shown that this species had the highest AR coverage in its leaf cuticle among Gramineae family. The significance of the project is to allow insights into a novel family of compounds in the plant cuticular wax through chemical analysis. Efforts were also made to clone the gene of alkylresorcinol synthase from tender rye (Secale cereale) leaves. However, those experiments made only limited progress and will therefore be described only briefly in Appendix B. The findings of chemical analysis are expected to provide a better underpinning for future biological and biochemical studies into rye wax ARs. Gene cloning and characterization is being continued in our lab.  13  References (1)  Jeffree, C. E. In Insects and The Plant Surface; Southwood, B. J. R., Ed.; E.  Arnold: London, 1986, p 23-135. (2)  Vogg, G.; Fischer, S.; Leide, J.; Emmanuel, E.; Jetter, R.; Levy, A. A.; Riederer,  M. J. Exp. Bot. 2004, 55, 1401-1410. (3)  Furstner, R.; Barthlott, W.; Neinhuis, C.; Walzel, P. Langmuir 2005, 21, 956-  (4)  Neinhuis, C.; Barthlott, W. Annals of Botany 1997, 79, 667-677.  (5)  Krauss, P.; Markstadter, C.; Riederer, M. Plant cell and Environ 1997, 20,  961.  1079-1085. (6)  Jacobs, J. F.; Koper, G. J. M.; Ursem, W. N. J. Progress in Organic Coatings  2007, 58, 166-171. (7)  Holmes, M. G.; Keiller, D. R. Plant Cell and Environ 2002, 25, 85-93.  (8)  Eigenbrode, S. D.; Jetter, R. Integr. Comp. Biol. 2002, 42, 1091-1099.  (9)  Goodwin, S. M.; Edwards, C. J.; Jenks, M. A.; Wood, K. V. Hortscience 2007,  42, 1631-1635. (10)  Eigenbrode, S. D.; Pillai, S. K. J. Chem. Ecol. 1998, 24, 1611-1627.  (11)  Eigenbrode, S. D.; Espelie, K. E. Annual Review of Entomology 1995, 40, 171-  (12)  Jenks, M. A.; Joly, R. J.; Peters, P. J.; Rich, P. J.; Axtell, J. D.; Ashworth, E. N.  194.  Plant Physiol. 1994, 105, 1239-1245. (13)  Kunst, L.; Samuels, A. L. Progress in Lipid Research 2003, 42, 51-80.  (14)  Markstadter, C.; Federle, W.; Jetter, R.; Riederer, M.; Holldobler, B.  Chemoecology 2000, 10, 33-40. (15)  Wen, M.; Buschhaus, C.; Jetter, R. Phytochem. 2006, 67, 1808-1817.  (16)  Xu, X. J.; Dietrich, C. R.; Delledonne, M.; Xia, Y. J.; Wen, T. J.; Robertson, D.  S.; Nikolau, B. J.; Schnable, P. S. Plant Physiol. 1997, 115, 501-510. (17)  Gable, K.; Garton, S.; Napier, J. A.; Dunn, T. M. J. Exp. Bot. 2004, 55, 543-545.  (18)  Beaudoin, F.; Wu, X. Z.; Li, F. L.; Haslam, R. P.; Markham, J. E.; Zheng, H. Q.;  Napier, J. A.; Kunst, L. Plant Physiol. 2009, 150, 1174-1191.  14  (19)  Bach, L.; Michaelson, L. V.; Haslam, R.; Bellec, Y.; Gissot, L.; Marion, J.; Da  Costa, M.; Boutin, J. P.; Miquel, M.; Tellier, F.; Domergue, F.; Markham, J. E.; Beaudoin, F.; Napier, J. A.; Faure, J. D. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 14727-14731. (20)  Vioque, J.; Kolattukudy, P. E. Arch. Biochem. Biophys 1997, 340, 64-72.  (21)  Rowland, O.; Zheng, H. Q.; Hepworth, S. R.; Lam, P.; Jetter, R.; Kunst, L. Plant  Physiol. 2006, 142, 866-877. (22)  Lardizabal, K. D.; Metz, J. G.; Sakamoto, T.; Hutton, W. C.; Pollard, M. R.;  Lassner, M. W. Plant Physiol. 2000, 122, 645-655. (23)  Li, F.; Wu, X.; Lam, P.; Bird, D.; Zheng, H.; Samuels, L.; Jetter, R.; Kunst, L.  Plant Physiol. 2008, 148, 97-107. (24)  Kozubek, A.; Tyman, J. H. P. Chem. Rev. 1999, 99, 1-25.  (25)  Ross, A. B.; Shepherd, M. J.; Schupphaus, M.; Sinclair, V.; Alfaro, B.; Kamal-  Eldin, A.; Aman, P. J. Agric. Food Chem. 2003, 51, 4111-4118. (26)  Ross, A. B.; Kamal-Eldin, A.; Jung, C.; Shepherd, M. J.; Aman, P. J. Sci. Food  Agric. 2001, 81, 1405-1411. (27)  Hengtrakul, P.; Lorenz, K.; Mathias, M. J. Food Compos. Anal. 1991, 4, 52.  (28)  Kozubek, A.; Demel, R. A. Biochem. Biophys. Acta 1980, 603, 220-227.  (29)  Chaturvedula, V. S. P.; Schilling, J. K.; Miller, J. S.; Andriantsiferana, R.;  Rasamison, V. E.; Kingston, D. G. I. J. Nat. Prod. 2002, 65, 1627-1632. (30)  Droby, S.; Prusky, D.; Jacoby, B.; Goldman, A. Physiol. Mol. Plant Pathol 1987,  30, 285-292. (31)  Diogenes, M. J. N.; deMorais, S. M.; Carvalho, F. F. Contact Dermatitis 1996,  35, 114-115. (32)  Garcia, S.; Garcia, C.; Heinzen, H.; Moyna, P. Phytochem. 1997, 44, 415-418.  (33)  Reffstrup, T.; Boll, P. M. Phytochem. 1985, 24, 2563-2565.  (34)  Montsant, A.; Zarka, A.; Boussiba, S. Marine Biotechnology 2001, 3, 515-521.  (35)  Magnucka, E. G.; Suzuki, Y.; Pietr, S. J.; Kozubek, A.; Zarnowski, R. Pesticide  Biochemistry and Physiology 2007, 88, 219-225. (36)  Streibl, M.; Konecny, K.; Trka, A.; Ubik, K.; Pazlar, M. Collection of  Czechoslovak Chemical Communications 1974, 39, 475-479. (37)  Tulloch, A. P.; Hoffman, L. L. Phytochem. 1974, 13, 2535-2540. 15  (38)  Dayan, F. E.; Kagan, I. A.; Rimando, A. M. J. Biol. Chem. 2003, 278, 28607-  (39)  Fate, G. D.; Lynn, D. G. J. Am. Chem. Soc. 1996, 118, 11369-11376.  (40)  Dimroth, P.; Ringelmann, E.; Lynen, F. Eur. J. Biochem. 1976, 68, 591-596.  (41)  Raharjo, T. J.; Chang, W. T.; Choi, Y. H.; Peltenburg-Looman, A. M. G.;  28611.  Verpoorte, R. Plant Sci. 2004, 166, 381-385. (42)  Suzuki, Y.; Kurano, M.; Esumi, Y.; Yamaguchi, I.; Doi, Y. Bioorg. Chem. 2003,  31, 437-452. (43)  Hesk, D.; Craig, R.; Mumma, R. O. J. Chem. Ecol. 1992, 18, 1349-1364.  16  Chapter 2: Alkylresorcinol deposition in the cuticle along rye (Secale cereale) leaves*  2.1 Introduction Based on the literature review in section 1.3.3, it is not clear whether ARs in those examples accumulated partly or entirely in the cuticular wax layer, and whether they are aimed at the cuticle to play biological functions at the plant surface. It is also not apparent how ARs might be distributed between the epi- and intracuticular wax layers. Preliminary experiments revealed relatively higher coverage of ARs on rye leaves than on leaves of other species in the Poaceae family. Therefore, rye (Secale cereale L. cv. Esprit) leaves were chosen for chemical analysis. The goal of this chapter was to address the questions (1) what compounds constitute the waxes of rye leaves and whether the leaf cuticular waxes contain ARs, (2) whether the abaxial and adaxial waxes have different wax composition, (3) whether ARs are restricted to the cuticular wax, and (4) whether the epicuticular wax layer is different from the intracuticular layer and where ARs are located throughout the two layers.  2.2 Materials and methods 2.2.1 Plant materials and growth conditions Rye (Secale cereale L. cv. Esprit) grains were obtained from Capers, Vancouver and planted in soil. Batches of plants were grown in plastic pots (diameter 15 cm) in a greenhouse at The University of British Columbia (23 °C, 14 h light at approximately 120 µmol m-2 s-1, relative humidity 75%). The second leaves were harvested before the third leaves had reached 5 cm long (typically three weeks after germinating). From 10 cm away from the point of emergence (POE) from the sheath of the first leaf, a 5 cm-long segment was cut out for wax analysis. Using one of three sampling techniques (see section 2.1.2), wax was extracted from the segment. Six independent parallels were analyzed for each technique. Surface areas of the sampled segments were calculated by multiplying leaf widths and segment length. *A version of this chapter has been published. Ji, X. and Jetter, R. (2008) Very Long Chain Alkylresorcinols Accumulate in the Intracuticular Wax of Rye (Secale cereale L.) Leaves near the Tissue Surface. Phytochem. 69:1197-1207.  17  2.2.2 Wax extraction Three plants were pooled for each sample. Six parallels were performed on independent samples rather than being just repeated GC runs of the same sample. In total, 18 plants were used for each extraction. The mixture of total wax was sampled by dipping the whole leaf segments twice for 30 s into CHCl3 containing n-tetracosane as the internal standard. The resulting extracts were concentrated, transferred into small GC vials, brought to dryness under a gentle stream of N2, and then stored at 4°C. Either adaxial or abaxial wax was selectively extracted using the brushing technique. Each side of the rye leaf segments was brushed 60 times with CHCl3-soaked fabric glass wool. Prior to use, the fabric glass wool was thoroughly delipidated with CHCl3 using a Soxhlet apparatus. After brushing, fabric glass was soaked in CHCl3 for half an hour to allow complete release of wax into CHCl3. Tetracosane was then added as the internal standard and the resulting solutions were dried and stored at 4 °C as described above for total wax. The epicuticular wax layer was discriminated from intracuticular wax by adhesive treatments using gum Arabic as an adhesive. Then solvent extraction was utilized to sample the intracuticular wax layer. Before use, gum Arabic powder (Sigma–Aldrich) was thoroughly extracted in CHCl3 using a Soxhlet apparatus, and dissolved in distilled water to make a glue solution of 1 g/mL. The adhesive solution was evenly spread onto the abaxial side of the leaf segment and air-dried. Approximately 40 min later, the hardened sticky film was peeled off and extracted with CHCl3 containing n-tetracosane as the internal standard. The process was repeated on the same side of the leaf segment until no significant wax could be extracted. The resulting solutions were combined, dried and stored at 4°C. After the mechanic removal of the outer layer, the same side of the leaf segment was brushed 20 times with fabric glass soaked with CHCl3 to extract the intracuticular wax. 2.2.3 Synthesis of 5-n-tridecylresorcinol (6, AR13:0) AR13:0 (6) was synthesized using the same method as described by Furstner and Seidel.1  18  2.2.4 Synthesis of 5-n-nonadecylresorcinol (1, AR19:0)  OCH3 (1)  O  F3C S O S  + H3CO  O  OH  O  7  CF3  O  H3C N CH3 CH2Cl2, 0 oC, 2h  OCH3 O H3CO  O-S-CF3 8  (2)  O  THF, Room temp., 2h  B-H +  B 9  (3)  8  +  9  THF, PdCl2(dppf), reflux, 1h  OCH3 H3CO 10  (4) B-I  +  10  OH  Hexane, room temp., 4h HO  1  Fig. 2.1 Synthesis of 1,3-dihydroxy-5-nonadecylbezene (AR19:0).  Synthesis of AR19:0 (1) was carried out using similar methods as described by Furstner and Seidel (Fig.2.1)1: triflic anhydride (2.1 g, 7.4 mmol, Sigma–Aldrich) was added to CH2Cl2 (9 ml) and the solution was slowly added to a mixture of 3,5-dimethoxyphenol (7) (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. The above mixture was cooled to 0°C and stirred for 2 h before adding 5 ml of H2O. The organic layer was separated and dried with Na2SO4. After evaporation of solvent under vacuum, the crude product was purified by flash column chromatography (CC) through packed silica gel with hexane:CH2Cl2 (1:1). Following solvent removal to dryness overnight, 3,5dimethoxyphenol triflate (8) was generated as a yellow syrup with yield of 90%. Next, the mixture  of  1-tridecene  (474  mg,  0.7  ml,  2.6  mmol,  Sigma–Aldrich)  and  9-  borabicyclo[3.3.1]nonane (9-BBN) (5.2 ml, 2.6 mmol, Sigma–Aldrich) in THF (60 ml) were stirred for 2 h under N2 at room temperature. Subsequently, NaOMe (0.17 g, 3 mmol), triflate (8) (0.65 g, 2.3 mmol) and PdCl2 (dppf) (56 mg, 0.07 mmol, Sigma–Aldrich) were added, the mixture was heated until reflux began, which was maintained for 1 h, the solvent was evaporated and CH2Cl2 (10 ml) was added, instead. By passage of the CH2Cl2 solution through a short column of silica, insoluble residues were removed. After solvent removal, the crude 19  product (9) was purified by flash CC with hexane/ EtOAc (15:1) as eluent, to form 1,3dimethoxy-5-nonadecylbenzene (10) as a colorless solid (54% yield). 10 (0.5 g, 1.2 mmol) and 9-iodo-9-BBN (0.41 ml, 2.52 mmol, Sigma–Aldrich) were mixed in hexane (25 ml) and stirred for 3 h at room temperature. Following solvent evaporation under vacuum, 15 ml of Et2O was added to dissolve the residue. To precipitate the 9-BBN ethanolamine adduct, ethanolamine (0.14 ml, 2.2 mmol, Sigma–Aldrich) in THF (1 ml) was added and the mixture was stirred for 3 h. The precipitate was removed with filter paper and the filtrate was brought to dryness. At last, flash CC was utilized to purify the crude product by using hexane/EtOAc (2:1) as eluent, producing analytically pure 1 (61% yield).  2.2.5 Sampling ARs from internal tissue of rye leaves ARs in the internal tissue of rye leaves were extracted based on the method described by Deszcz and Kozubek.2 After removal of total surface wax by soaking leaves in CHCl3 twice for 30 s, rye leaves were homogenized in liquid N2 and immersed into CHCl3:MeOH (2:1, v/v) containing synthetic AR13:0 (6) as the internal standard. The resulting solution was concentrated and separated on TLC (silica) with CHCl3:EtOAc (85:15). The relevant bands were scraped off and soaked in CHCl3, the solvent was evaporated and the resulting extracts were stored at 4°C for analysis.  2.2.6 Chemical analysis Wax extracts were derivatized with N,O-bis(trimethylsilyl)-trifluoroacetamide (BSTFA; 10 µl Sigma–Aldrich) in pyridine (10 µL) at 70°C for approximately 30 min in order to transform all hydroxyl groups into the corresponding trimethylsilyl (TMSi) ethers, for example in Fig.2.2.3 The resulting solutions were diluted with 100 µL of CHCl3 before analysis by GC– MS and GC–FID. The wax mixtures were separated using capillary GC (5890 N, Agilent, Avondale, PA; column 30 m HP-1, 0.32 mm i.d., Agilent) with He carrier gas inlet pressure programmed for constant flow of 1.4 ml min-1 and the compositions were monitored by MS detector (Electron impact ionization, Acceleration voltage 70 eV, Quadrupole ion filter, scan range m/z 50 - 650, scan rate 0.3 s-1, 5973N, Agilent). Samples were on-column injected at 50°C and GC oven was temperature-programmed: 2 min at 50°C, increased to 200°C by 40°C min-1, held at 200°C for 20  2 min, raised to 320°C by 3°C min-1 and held at 320°C for 30 min. Identification of individual compounds was carried out with GC-MS by comparing characteristic fragments with those of authentic standards and data listed in literature. Quantification of individual compounds was performed against the internal standard by integrating peak areas using GC–FID under the same GC conditions as above, but with H2 carrier gas inlet pressure at a constant flow of 2 ml min-1. All quantitative data were presented as the means of six parallel experiments and standard deviations. SPSS 13.0 (SPSS, USA) was used for statistical analyses. F3C OH  HO  Si N  R  O Si  Pyridine, ∆  O  Si  O  Si  R  R=n-C19H39, n-C21H43, n-C23H47, n-C25H51, n-C27H55  Fig. 2.2 Derivatization reaction with N,O-bis(trimethylsilyl)-trifluoroacetamide.  21  2.3 Results and discussion Approximately 5-cm-long segments located about 10 cm above the point of emergence of the second leaves out of the sheath of first leaves were analyzed in order to minimize biological variation. Preliminary results indicated that wax amounts were relatively constant over time since the differentiation and growth of epidermal cells had stopped in that area. The chemical composition of rye leaf cuticular wax was carried out in three sets of experiments aimed at revealing the wax composition with varying levels of spatial resolution and to allow comparisons with the literature data. In a first experiment, the combined wax from both sides of the leaf was extracted, in order to get overall data of total wax compositions. A second experiment was designed to distinguish the composition of wax on abaxial and adaxial sides of rye leaves. A third chemical analysis was aimed at differentiating the wax composition from epi- and intracuticular wax layers.  2.3.1 Total wax In the first experiment, the combined leaf wax from both sides was extracted by brief immersion of the leaf segment in CHCl3. The overall wax load was 12.2 ± 1.5 µg/cm2 (Fig. 2.3), containing primary alcohols (71%), alkyl esters (11%), aldehydes (5%), alkanes (3%), steroids (0.3%), together with traces of secondary alcohols as well as fatty acids. Complete homologous series of fatty acids (C20-C34), primary alcohols (C22-C30) and aldehydes (C26–C32) were revealed (Table 2.1).4 They were all dominated by even-numbered homologs with C26 homologs strongly prominent in all three homologous series (Fig.2.4). The dominating constituent was hexacosanol, accounting for 69% (8.7 ±1.3 µg/cm-2) of the total wax mixture. The chain length distribution of alkanes, ranging from C27 to C33 (Table 2.1), was dominated by odd-numbered homologs and showed a maximum at C31 (Fig. 2.4). The alkyl ester fraction consisted of even-numbered homologs ranging from C40 to C52, and had a maximum at C44 (Table 2.2).4  22  2  Wax Load (µg/cm )  10  Total Adaxial Abaxial  8  6  4  2  Unidentified  Steroids  Alkylresorcinols  Secondary alcohols  Alkanes  Aldehydes  Alkyl esters  Primary alcohols  Fatty acids  0  Fig. 2.3 Compound class composition of cuticular wax mixtures from the second leaf of rye. The coverages of all compound classes are given as means (n=6) and SD for the total wax extracted from both sides of the leaf together, and for the wax extracted from the adaxial and abaxial sides separately.  Comparing with literature data, my findings are similar in wax composition with other reported Poaceae species. Alkanes, primary alcohols, fatty acids and aldehydes were consistently found in rice (Oryza sativa), wheat (Triticum aestivum and T. durum), barley (Hordeum vulgare) and maize (Zea mays) waxes as well as on rye leaves.5-7  23  Table 2.1 Very long chain compounds identified in rye leaf waxes. 4  Note: Wax mixtures were sampled by extracting both sides of the leaf together (To), by extracting the adaxial (Ad) or the abaxial (Ab) sides selectively, or by removing the epicuticular (Ep) and the intracuticular (In) wax layers consecutively from the abaxial side. Predominant homologs are highlighted in bold face.  Table 2.2 Homolog and isomer composition of very long chain alkyl esters identified in rye leaf waxes.4  Note: Wax mixtures were sampled by extracting both sides of the leaf together (To), by extracting the adaxial (Ad) or the abaxial (Ab) sides selectively, or by removing the epicuticular  24  (Ep) and the intracuticular (In) wax layers consecutively from the abaxial side. Predominant acyl homologs are highlighted in bold face.  Fig. 2.4 Chain length distributions of individual wax compounds in the total, adaxial and abaxial wax mixtures on the second leaf of rye. Percentages of individual homologs within the series of (A) fatty acids, (B) primary alcohols, (C) aldehydes and (D) alkanes are shown as means (n=6) with SD.  For rye cuticular wax, similar detail has not been studied to date. Besides those common wax components, the wax mixture extracted from rye straw contained high amounts of βdiketones not found in this study.8 Very similar findings were published for the wax sampled from all leaves together.9 This discrepancy between our findings and the literature data might be attributed to not only variability between rye cultivars, but more importantly to various organs from which wax mixtures were extracted. Since β-diketones were found to be the dominating wax components from barley spikes and stem,10,11 and from wheat flag leaves,12 it can be inferred that β25  diketones reported in rye straw wax were likely to stem from those organs rather than lower leaves studied in present chapter.  2.3.2 Identification of alkylresorcinols in cuticular wax OH  HO  R*  R  Name  Number in text  Abbreviation  C19H39  5-Nonadecylresorcinol  1  AR19:0  C21H43  5-Heneicosylresorcinol  2  AR21:0  C23H47  5-Tricosylresorcinol  3  AR23:0  C25H51  5-Pentacosylresorcinol  4  AR25:0  C27H55  5-Heptacosylresorcinol  5  AR27:0  C13H27  5-Tridecylresorcinol  6  AR13:0  * R are n-alkyl chains.  Fig. 2.5 Structures of alkylresorcinols identified in rye eaf wax (1-5) and have synthesized as standards for structure elucidation (1) as well as quantification (6).4  Besides those typical wax components mentioned above, there were five even-spaced compounds that had not been reported before from any leaf cuticular wax. Their molecular ions differed by 28 mass units, suggesting a homologous series differing by –CH2CH2- units. All of them showed characteristic peaks of alkylresorcinol TMSi ethers at m/z 73, 268 and 281,13,14 and their corresponding fragments [M-15]+ indicated the loss of a methyl group from the TMSi derivatives. To confirm that the five compounds are ARs, one representative (AR19:0) of the homologous series was chemically synthesized and run in GC-MS for comparison. The AR19:0 standard showed identical mass spectrum with one of the unidentified homlogs. Moreover, they co-eluted under the GC conditions used. This result not only verified the structure of this wax AR, but also excluded other isomers. It had been reported that GC could separate various 26  isomers of comparable phenolics, including isomers with both side-chain and ring-positional configuration, due to their different physical properties.15 Hence, individual AR isomers could usually be identified by joint analysis using GC-MS even though they may not be discriminated by MS alone. Therefore, the comparison with an authentic standard using GC-MS unambiguously proved the structure of one of the five compounds to be AR 19:0 (1).  50 Total Adaxial Abaxial  Percentages [%]  40 30 20 10 0 19  21 23 25 Alkyl chain length  27  Fig. 2.6 Chain length distributions of alkylresorcinols in the total, adaxial and abaxial wax mixtures on the second leaf of rye. Percentages of individual homologs within the fraction are shown as means (n =6) and SD.  All the evidence taken together, the five wax constituents were identified as a homologous series of ARs with alkyl chain lengths C19, C21, C23, C25, and C27 (1-5) (Table 2.1, Fig. 2.5).4 Homologs with odd-numbered alkyl side chains dominated the series, especially AR 21:0 (2), 23:0 (3) and 25:0 (4) (Fig. 2.6). ARs as a compound class had a coverage of 0.3 ± 0.1 µg/cm2 on rye leaves, corresponding to approximately 3% of the total wax. Only 2 % of total wax remained unidentified after the identification of ARs.  27  2.3.3 Alkylresorcinols in internal tissue To test whether ARs are restricted to the cuticular wax or whether they also exist in internal tissue of rye leaves, internal ARs were extracted after surface waxes had been initially removed. With synthetic AR13:0 as internal standard, ARs were quantified using the intensity of their characteristic peak (m/z 268) in mass spectra. Internal ARs were discovered to be 9 µg/g, contrasted to 135 µg/g of cuticular ARs. Hence, cuticular ARs contributed about 94% of the total ARs of rye leaves. ARs were not mentioned in previously published papers about rye waxes.8,9 Yet, they were reported in total lipid extracts from various organs of rye. In rye grains, Montsant et al.16 reported 559 µg/g dry weight of ARs ranging from AR 17:0 to 25:0 (odd numbers only), with AR 17:0, 19:0 and 21:0 as the main homologs.16 Rye seedlings were found to contain 3.1 µg/g dry weight of ARs from AR15:0 to 25:0 and a strong predominance of AR17:0, AR19:0 and AR23:0.17 Unfortunately, it is not clear whether the ARs were accumulated in the cuticular wax of those organs or whether they were located exclusively or partially in the internal tissues. Further investigations should be performed in those organs to differentiate between these possibilities. One interesting finding about the alkyl chain lengths of ARs in either rye grains or seedlings is the lower homologs than cuticular ARs of rye leaves. It was also found that leaf cuticular ARs peak at longer chain lengths than those of other organs among their homologuous series.  2.3.4 Abaxial and adaxial wax To test whether ARs were restricted to one side of rye leaves, the second chemical analysis was executed to differentiate between the waxes from the abaxial and adaxial surfaces of S. cereale leaves. The second method was established for selective extraction of waxes from either side of gymnosperm needles.18 The leaf was gently brushed with fabric glass pre-soaked with CHCl3, in order to completely remove the treated surface while keeping intact the opposite side of the leaf. Initially, rye leaf surfaces were brushed 20 times, producing only two thirds of total wax yields. Yet, when brushed more than 60 times, the extracts began to appear green, indicating contamination by chlorophyll and indicated that the boundary of internal tissues was reached. Thus, 60 times brushing was used as a standard protocol to selectively extract waxes on either side. The yields were 12.1±2.5 µg/cm2 and 11.5±2.0 µg/cm2 from the adaxial and 28  abaxial surfaces, respectively. Comparing with the total wax coverage found in the first experiment, the overall average wax load (11.8 ± 3.2 µg/cm2) of both sides was not significantly different. This finding confirmed that the brushing technique allowed selective and exhaustive extraction of adaxial and adaxial waxes. Both adaxial and abaxial waxes had all the compound classes previously identified in total wax from both sides combined. Overall, no significant difference was found either between the compound class percentages on both sides or between the homolog patterns within these classes (Tables 1 and 2, Fig. 2.4). ARs were discovered to have approximately equal load of 0.2 ±0.1 µg/cm2 on the adaxial side (contributing 1.4 ± 0.4% of total adaxial wax) and of 0.3 ±0.1 µg/cm2 on the abaxial side (contributing 2.2 ± 0.5% of abaxial wax). Similarly, the homolog patterns in the AR fractions from both sides of the leaves were found to resemble each other (Table 2.1, Fig. 2.6). Overall, the results manifest that the adaxial and abaxial wax covered on rye leaves share relatively similar wax composition. Hence, rye leaf can serve as an example where two surfaces of the same organ can share the same overall makeup of cuticular wax, even though the two surfaces have different physiological and ecological significance. Relatively few other species had been analyzed for adaxial and abaxial waxes separately. And in many cases, adaxial and abaxial wax exhibited differences in composition. For example, the adaxial wax of Pisum sativum contained predominantly primary alcohols, whereas the corresponding abaxial wax was found to have large amounts of alkanes.19-21  2.3.5 Epi- and intracuticular wax The third experiment was performed to examine whether compositional differences occurred between epi- and intracuticular wax layer of S. cereale leaves. This was also to address the question whether ARs were restricted in the inner wax layer or exposed at the leaf surface, or whether they existed all the way through the leaf cuticular wax. Since no obvious differences had been marked between the compositions of the adaxial and abaxial waxes, this last experiment focused merely on the abaxial sides of the leaves, from which side the epi- and intrcuticular waxes were to be selectively sampled.  29  80 Epicuticular Intracuticular Total abaxial  Percentages [%]  70 60 50 40 30 20 10  Unidentified  Steroids  Alkylresorcinols  Secondary alcohols  Alkanes  Aldehydes  Alkyl esters  Primary alcohols  Fatty acids  0  Fig. 2.7 Relative amounts of wax compound classes within the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye. The results from the four gum arabic treatments were taken together to show the composition of the epicuticular waxes, whereas the final extraction results represent the composition of the intracuticular waxes.  Mechanical removal of surface wax layer was performed with Gum Arabic as the glue. The same leaf could be treated four consecutive times without damaging the abaxial surface. The gum arabic treatments could not be repeated more often because leaves were too fragile. The successive adhesive treatments yielded slightly decreasing wax amounts of 2.3± 1.0, 2.3 ± 0.5, 2.2 ±0.5, 2 and 1.7 ±0.4 µg/cm-2. Statistical comparison between the cumulative wax yields after the 3rd and 4th treatments resulted in no significant difference (ANOVA, N=6, P =0.066), suggesting that the four consecutive treatments together had thoroughly sampled the 30  mechanically accessible wax and a fifth glue treatment would not have added more material. Similarly, it can be concluded that sampling in this way had reached the mechanically resistant cutin matrix and therefore had removed epicuticular wax exhaustively. After exhaustive removal of epicuticular wax, the leaf was finally extracted with organic solvent. The corresponding wax yield after extracting twice with CHCl3 (6.0 ± 2.0 µg/cm-2)  was  significantly different from the yield of the 4th mechanical removal (ANOVA, N=6, P<0.0005), indicating that the final CHCl3 extraction sampled wax from a distinct layer inside the cuticle. Corresponding wax was inferred to be intracuticular wax. The combined wax harvest in this experiment including epi- and intracuticular wax matched closely the yield of adaxial wax in the second experiment (ANOVA, N =6, P = 0.160), confirming the efficience of the sampling methods. 100  A  Epicuticular Intracuticular Total abaxial  Percentages [%]  80 60 40  Percentages [%]  100  20  60 40  0 24  26 28 Chain length  30  24  C  30  80 Percentages [%]  Percentages [%]  80  20  0  100  B  60 40  26 28 Chain length  30  D  20  10  20 0  0 26  28 30 Chain length  32  27  28  29 30 31 Chain length  32  33  Fig.2.8 Chain length distributions of individual wax components in the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye. Percentages of individual homologs within the series of (A) fatty acids, (B) primary alcohols, (C) aldehydes and (D) alkanes are shown as means (n=6) with SD. 31  Overall, the wax composition was homogeneous throughout the epi- and intracuticular wax layers. The two wax layers shared very similar composition and relative amounts, both in terms of compound classes and chain length distributions (Tables 2.1 and 2.2, Figs. 2.7 and 2.8). One of the two exceptions were the steroids, showing higher wax coverage and percentage in the intracuticular layer than the epicuticular layer. The result corresponded to the previous findings in other species, where cyclic wax constituents displayed gradients from higher concentration inside the cuticle to lower concentration on the surface.18 The most drastic gradient between the two wax layers of rye leaves was revealed for the ARs. ARs could not be detected in epicuticular layer and were confined entirely to the intracuticular wax layer (Fig. 2.7). AR concentration in the latter layer was 0.2 ± 0.1 µg/cm2, contributing 2.0 ± 0.8% of the total intracuticular wax. Overall, intracuticular ARs closely matched the overall adaxial ARs in terms of qualitative and quantitative chain length distributions (Fig. 2.9). All the results taken together, it can be concluded that ARs found in the rye leaf cuticle are restricted to the intracuticular wax layer.  50 Epicuticular Intracuticular Total abaxial  Percentages [%]  40 30 20 10 0 19  21 23 25 Alkyl chain length  27  Fig. 2.9 Chain length distributions of alkylresorcinols in the epicuticular, intracuticular, and total abaxial wax mixtures on the second leaf of rye. Percentages of individual homologs within the fraction are shown as means (n= 6) with SD. 32  2.4 Conclusion To sum up, ARs with alkyl side chain lengths ranging from C19 to C27 were discovered in the leaf cuticular wax of rye. The leaf ARs were restricted largely to the cuticle, with quite low concentrations found in the internal tissue. Both sides of rye leaves exhibited similar amounts of cuticular ARs, while only intracuticular wax layer contained ARs and epicuticular wax layer was devoid of them. This finding indicates that rye leaf ARs are covered by a thin epicuticular wax layer even though they are placed fairly close to the very surface of rye leaves. It can be concluded that the majority of ARs are not located at the outer surface of rye leaves and, thus, are not accessible for direct interaction with microorganisms or insects that come into direct contact with the leaf surface.  33  References (1)  Furstner, A.; Seidel, G. J. Org. Chem. 1997, 62, 2332-2336.  (2)  Deszcz, L.; Kozubek, A. Biochimica Et Biophysica Acta-Molecular and Cell  Biology of Lipids 2000, 1483, 241-250. (3)  Deas, A. H. B.; Baker, E. A.; Holloway, P. J. Phytochem. 1974, 13, 1901-1905.  (4)  Ji, X. F.; Jetter, R. Phytochem. 2008, 69, 1197-1207.  (5)  Tulloch, A. P.; Hoffman, L. L. Phytochem. 1971, 10, 871-&.  (6)  Bianchi, G.; Lupotto, E.; Russo, S. Experientia 1979, 35, 1417-1417.  (7)  Reynhardt, E. C.; Riederer, M. European Biophysics Journal with Biophysics  Letters 1994, 23, 59-70. (8)  Streibl, M.; Konecny, K.; Trka, A.; Ubik, K.; Pazlar, M. Collection of  Czechoslovak Chemical Communications 1974, 39, 475-479. (9)  Tulloch, A. P.; Hoffman, L. L. Phytochem. 1974, 13, 2535-2540.  (10)  Wettstein-Knowles, v. In Biochemistry and Metabolism of Plant Lipids; Kuiper,  J. F. G. M. W. P. J. C., Ed.; Elsevier: 1982, p 69-78. (11)  Richardson, A.; Wojciechowski, T.; Franke, R.; Schreiber, L.; Kerstiens, G.;  Jarvis, M.; Fricke, W. Planta 2007, 225, 1471-1481. (12)  Tulloch, A. P. Phytochem. 1973, 12, 2225-2232.  (13)  Linko, A. M.; Parikka, K.; Wahala, K.; Adlercreutz, H. Analytical Biochemistry  2002, 308, 307-313. (14)  Ross, A. B.; Shepherd, M. J.; Schupphaus, M.; Sinclair, V.; Alfaro, B.; Kamal-  Eldin, A.; Aman, P. J. Agric. Food Chem. 2003, 51, 4111-4118. (15)  Fritz, J. O.; Moore, K. J. J. Agric. Food Chem. 1987, 35, 710-713.  (16)  Montsant, A.; Zarka, A.; Boussiba, S. Marine Biotechnology 2001, 3, 515-521.  (17)  Magnucka, E. G.; Suzuki, Y.; Pietr, S. J.; Kozubek, A.; Zarnowski, R. Pesticide  Biochemistry and Physiology 2007, 88, 219-225. (18)  Wen, M.; Buschhaus, C.; Jetter, R. Phytochem. 2006, 67, 1808-1817.  (19)  Holloway, P. J.; Hunt, G. M.; Baker, E. A.; Macey, M. J. K. Chemistry and  Physics of Lipids 1977, 20, 141-155. (20)  Simon, S. D. E. C. W. M. R. C. J. Pisum Genet. 1998, 20 13–17.  34  (21)  Gniwotta, F.; Vogg, G.; Gartmann, V.; Carver, T. L. W.; Riederer, M.; Jetter, R.  Plant Physiol. 2005, 139, 519-530.  35  Chapter 3: Time course of alkylresorcinol deposition in the cuticle along the rye (Secale cereale L.) leaf*  3.1 Introduction In chapter 2, it was shown that ARs accumulate in the cuticular waxes on rye leaves and that they are limited to the intracuticular wax layer. This leaves the question whether these compounds are found in the cuticular wax mixture due to accidental formation as a side product of fatty acid biosynthesis in epidermal cells and due to passive lipophilic partitioning, or whether they are the product of a dedicated biosynthetic machinery. In order to distinguish between these possibilities, the genes and enzymes involved in their formation must be characterized. To this end, the relevant genes must be cloned using mRNA harvested from ARproducing cells, and the latter can be localized by detailed chemical studies of leaves in various developing stages and locations along the growing leaf. Monocotyledonous plants, such as grasses, are fundamentally different from dicotyledonous plants in terms of leaf expansion and cuticular wax accumulation.1 In dicotyledonous plants, growth of leaf area is due to leaf expansion in both longitudinal and lateral directions, leaving dicot leaves their characteristic broad-shaped look. Cuticular waxes must be deposited incessantly to match leaf expansion. In contrast, in monocot plants, increase of leaf surface area is mainly caused by longitudinal expansion of cells, in many grass species within sheaths of older leaves. When cells exit the elongation zone which is 20-60 mm from the base of leaf, they continue some lateral expansion before appearing from the enclosed sheath into sight. At that time period, cuticular waxes must be adequately biosynthesized and placed onto surface to fulfill their functions.1 In dicots, the time course of cuticular wax accumulation together with leaf development had been extensively studied.2,3 However, for monocots, only two studies for leek (Allium porrum) and barley (Hordeum vulgare) were reported. 1,4 Using hexacoanol as the representative of cuticular wax, Richardson et al. found that the coverage of wax along barley leaf was related to the distance from the point of emergence (POE) of the third leaf out of the *A version of this chapter will be submitted for publication. Ji, X. and Jetter, R. (2010) Time Course of Alkylresorcinol Deposition in the Cuticle along the Rye (Secale cereale L.) leaves.  36  sheath of the second leaf.1,5 Unfortunately, they only monitored until the time point when the targeted leaf was 3 cm long, missing the later stages. In addition, the approach using hexacosanol as the representative of overall cuticular wax ignored important information on other wax constituents, such as secondary alcohols, alkanes, alkyl esters as well as alkylresorcinols (ARs).  Considering the limited information on the time course for  monocotyledonous plants, more plant species needed to be investigated. Therefore, the goals of this chapter were to study the total cuticular wax over an extended period of time: (1) to monitor the wax composition over time, (2) to investigate the distribution of wax along the leaf over time, (3) to examine wax accumulation on the same segment over time, and (4) more importantly, to investigate the distribution and accumulation of ARs along the leaf over time. Rye (Secale cereale L.) was chosen as the model plant for the study. Preliminary studies had shown that there are no ARs detectable in the wax before the POE. Thus, only the segments beyond the POE, and time points during the development of the second leaf past the POE were studied.  3.2 Materials and methods 3.2.1 Plant materials and growth conditions Rye (Secale cereale L. cv. Esprit) seeds were purchased from Capers, Vancouver and germinated directly in soil. Plants were grown in several batches in plastic pots (diameter 15cm) at a density of 18-22 plants per pot in a growth chamber at The University of British Columbia (20°C, 24 h continuous light at 90-120 µmol m-2 s-1, relative humidity 70% ). Leaf two emerged from the sheath of leaf one at day 5 after germination. Starting at that point, growth of leaf two was monitored daily by measuring the length of the blade beyond the point of emergence (POE) with a ruler. For analysis, the second leaves of three plants each were harvested at growth stages I (7 cm long, plant 7 - 8 d old), II (10 cm long, plant 9 - 11 d old), III (15 cm long, plant 12 - 14 d old) and IV (20 cm long, plant 15 - 17 d old) (Fig. 3.1). Only those with exact lengths of 7, 10, 15 and 20 cm were used for chemical analysis. Leaves harvested at stages I – III were cut into 1 cm-segments, while those from stage IV were cut into 2 cm-segments. Corresponding segments from three different leaves were pooled together as one sample for wax extraction. Six 37  independent parallels were analyzed for each position at each growth stage. Extracted surface areas were calculated based on leaf widths and segment length, and accounting for both upper and lower sides of the leaves. Leaf two Leaf one  Point of Emergence (POE)  Sheath  a  Growth stage IV (20cm, 15-17 d) Growth stage III (15cm, 12-14 d)  20 18 16  Growth stage II (10cm, 9-11 d) Growth stage I (7cm, 7-8 d) 20 19 18 17 16 15 14  b  20 19 18 17 16 15 14 13 12 11  20 19 18 17 16 15 14 13 12 11 10 9 8 7 6  c  14 12 10 8 6 4 2  d  e  Fig. 3.1 Schematic overview of the sampling design employed in this study. The length of leaf two was measured from the point of emergence (POE) out of the sheath of leaf one upward to its end (a). Leaves were harvested at four different growth stages (b-e), and cut into 1 cm- or 2 cm-segments. Segments were numbered beginning at the leaf tip from 20 downwards in order to have consistent designations for corresponding segments between growth stages, and have the numbering sequence parallel segment age.  38  3.2.2 Wax extraction Prior to the extraction, known amounts of n-tetracosane and 5-n-tridecylresorcinol (AR13:0) were added as an internal standards to the solvent. Each plant sample was entirely submerged twice for 30 s into CHCl3 at room temperature. The resulting solutions were concentrated, transferred into small vials, brought to dryness under a gentle stream of nitrogen, and stored at 4°C. Three plants were pooled for each sample. Six parallels were performed on independent samples rather than being just repeated GC runs of the same sample. In total, 18 plants were used for each extraction.  3.2.3 Chemical analysis Wax extracts were derivatized with N,O-bis(trimethylsilyl)-trifluoroacetamide (BSTFA) under the same conditions as in Section 2.1.3. The resulting wax derivatives were diluted with CHCl3 prior to analysis by GC-MS and GC-FID under the same conditions as described in Section 2.1.4. The qualitative composition of the wax mixtures was studied using GC-MS. Individual compounds were identified by comparison of characteristic fragments with those of authentic standards and literature data. Homologs of alkylresorcinols were quantified against one of the internal standards, AR13:0, based on relative abundance of the characteristic fragment (m/z 268) in GC-MS runs. Other individual compounds were quantified against the other internal standard (n-tetracosane) by automatically integrating peak areas in GC-FID runs. All quantitative data are given as means of six parallel experiments and standard errors. Statistical analyses were performed with PAST software (PAST, USA). In order to determine the turning points between accumulation periods and steady concentration periods, nonlinear curves of the form y=a(1-e-bx) were fitted to the original data points, the beginning and final slopes of the curves were determined, and the point of intersection of the two tangent lines was thus regarded as the turning point.  39  3.3 Results and discussion The goal of this chapter was to monitor cuticular wax over the course of leaf development in order to find out when and where ARs deposit on the plant surface. Preliminary results showed that rye leaf one had very small amounts of ARs while leaf two, three and four had similar AR coverage. To allow quick supply of plant materials while reducing variability, only leaf two was chosen for the time course experiments. Since previous studies on a comparable species1 indicated that wax started to accumulate only near POE, in addition that preliminary investigation showed no AR in wax until several centimeters beyond the POE, the current analyses was limited on the parts of the leaf blade beyond the POE. First, the growth of leaf two was monitored by measuring length and width of the exposed leaf blade as a function of time, and then the development of cuticular wax on the exposed leaf blade was studied.  3.3.1 Leaf growth over time It was observed that since the leaf grew beyond the POE, it expanded mainly in length (Fig.3.2) yet rarely in width. This result coincides with findings by Kavanova et al.6 Hence leaf size could be assessed based on measurements of its length. Leaf length was measured starting from POE to the end of leaf two. The length increased linearly from 2.5 cm at day 6 to 20 cm at day 16. Growth rate was approximately 1.8 cm per day from day 6 to day 16 before the length reached about 21 cm. In order to assess wax development as a function of leaf development, four different time points (growth stages) were selected so that cuticular wax composition could be compared between them. The developmental stages at days 7, 10, 13 and 16 when the leaf length was about 7, 10, 15 and 20 cm long, were chosen as growth stage I, II, III and IV, respectively. The time intervals of the four stages were almost equal, with about three-day difference. At the four growth stages, only the leaves with exact length of 7, 10, 15 and 20 cm long respectively were harvested. Leaves harvested at each of these time points were cut into segments along their length axis, and the surface wax from each segment was investigated independently.  40  Length [cm]  20  15  10  5  0 6  7  8  9  10  11  12  13  14  15  16  17  18  19  20  Time [days after germination]  Fig. 3.2 Growth of the second leaf of rye. The length of the leaf blade beyond the point of emergence from the sheath was monitored as a function of time (measured as the number of days after germination). Data are given as averages from ten parallels with standard deviations.  41  3.3.2 Total waxes 3.3.2.1 Spatial distribution of total waxes At growth stage I (7 d old), the total amounts of cuticular wax were found to vary from 6 µg/cm2 at the POE (segment 14) to 9 µg/cm2 at the leaf tip (segment 20) (Fig.3.3). A steady increase of total wax amounts along the leaf axis was observed for the first 2 cm away from the POE (segments 14 – 16), whereas the rest of the leaf blade up to its tip showed constant wax loads. Thus, the spatial distribution of wax along the leaf axis indicated that wax biosynthesis in young leaves occurred at/near the POE, but not on the leaf blade beyond. Similar trends were observed for the later growth stages: waxes started with comparable wax loads at POE (6-8 µg/cm2), steadily increased for the first 2 cm, and then reached a plateau at uniformly wax amounts (9-10 µg/cm2) (Fig. 3.3). This indicated a 2-cm-long zone of active wax biosynthesis at/near POE was maintained throughout leaf development.  10.0  2  Wax Loads [µ µg/cm ]  12.0  8.0 6.0 4.0  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  2.0 0.0 0  2  4  6  8  10  12  14  16  18  20  Segment Position along the leaf Fig. 3.3 Distribution of total wax amounts along the second leaf of rye at four growth stages. The total wax coverages are given for each 1 cm-segment or 2 cm-segment starting from the point of emergence (POE) as averages of six parallels with standard deviations.  42  However, since the leaf was during the same time continuously growing, presumably by intercalation at its base as is characteristic for all monocotyledonous species, the wax accumulation zone could not consist of the same cells and tissue throughout that time. Instead, biosynthetically active (epidermis) cells must have been pushed into this zone at the POE and moved out of it at the distal end. Based on the result that the leaf was expanding at a rate of 1.8 cm/day, it can be concluded that the biosynthetic machinery in these cells was active for slightly more than one day after emergence. In this time period the leaf surfaces accumulated 2 - 3 µg/cm2 of wax, thus defining the rate of wax biosynthesis.  3.3.2.2 Accumulation of total waxes over time It was found that the wax loads on segments near/at POE increased considerably. For instance, the wax loads of segment 14 increased from 6 µg/cm2 to 9 µg/cm2 between growth stages I and II; segments 6 and 10 also accumulated an additional 2 - 4 µg/cm2 from one growth stage to the next. Roughly 3 µg/cm2 of wax was accumulated between two neighboring growth stages. Since there were only 3 d intervals between two adjacent stages, a wax accumulation rate at/near POE of approximately 1 µg/cm2 per day can be inferred. This is the accumulation rate inferred independently from the temporal pattern. Although this value is lower than the one inferred above from the spatial pattern (2-3 µg/cm2), they are within the same order of magnitude. Both results together give an impression of the rate of wax accumulation on the rye leaf beyond the POE.  43  3.3.3 Compound classes As discovered in Chapter II, the compound classes on rye leaves consisted of primary alcohols, secondary alcohols, alkyl esters, alkanes, fatty acids, aldehydes and alkylresorcinols. Primary alcohols were largely predominating (71.6-80.1% of total wax amounts). Alkyl esters and aldehydes were second and third, respectively, to primary alcohols while alkanes, fatty acids and ARs contributed less than 5% of the total wax. Altogether the seven compound classes comprised over 90% of the bulk wax at all times and locations. The compound classes of VLCFA derivatives shared a similar distribution pattern along the leaf except alkanes. Primary alcohols had about 5 µg/cm2 of coverage at the POE in all four growth stages, steadily increased from POE to 2 cm segments beyond POE, and reached a plateau of 7 - 8 µg/cm2 on the distal parts of the leaf blade (Fig. 3.4A). Alkyl esters, aldehydes and secondary alcohols followed similar trends with the following minor differences: (1) these compounds accumulated in zones that extended slightly farther away from the POE (2 - 4 cm), (2) the compounds had lower overall concentrations in the distal parts of the leaf, and (3) they showed more gradual transitions between the wax accumulation zone and plateau (Fig. 3.4B, C and E). The alkanes displayed a distinct spatial distribution pattern from the general trend. They were present at constant levels on all segments of the leaf (Fig. 3.4D). Despite all the small differences between spatial patterns for various compound classes, their relative portions did not shift significantly between leaf segments. For example, primary alcohols accounted for 71.6-77.8% of total wax loads on the segments within the wax production zone and for similar percentages, 77.5-80.1%, on other segments. There was also no change in the relative composition of compound classes over time, as seen in comparisons of the wax mixtures on the same leaf segment at various developmental stages (Fig. 3.5). Thus, the conclusions drawn above for the zones and time periods in which bulk wax biosynthesis occurs also apply to the formation of all the major compound classes. ARs were formed in a remarkably different spatial area and in distinct time periods. They were constantly detected at growth stages III and IV. At growth stage IV, ARs were not detectable at the POE. ARs started to show up at segment 4 (4 cm away from POE) and the amounts peaked at 0.2-0.3 µg/cm2 in segment 4-10, i.e. 2 – 10 cm away from the POE (Fig. 3.4F). Then the further distal portion of the leaf blade (10 – 20 cm away from the POE) was covered by small amounts of ARs at approximately 0.1 µg/cm2. ARs were present at relatively 44  1.6  A  1.4  8.0 6.0 4.0  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  2.0  Alkyl ester 2 loads [µ µg/cm ]  prim. Alcohol 2 loads [µ µg/cm ]  10.0  1.2 1.0 0.8 0.6 0.4 0.2  0.0  0.0 0 2 4 6 8 10 12 14 16 18 20  0 2 4 6 8 10 12 14 16 18 20 Segment position along the leaf  Segment position along the leaf  C  0.6 Alkane 2 loads [µ µg/cm ]  Aldehyde 2 loads [µ µg/cm ]  0.6  0.4  0.2  0.4  0.2  0 2 4 6 8 10 12 14 16 18 20 Segment position along the leaf  0 2 4 6 8 10 12 14 16 18 20 Segment position along the leaf  0.4  F  E Alkylresorcinol loads [µ µg/cm2]  sec. Alcohol  loads [µ µg/cm2]  0.12  D  0.0  0.0  0.14  B  0.10 0.08 0.06 0.04  0.3 0.2 0.1  0.02 0.0  0.00 0 2 4 6 8 10 12 14 16 18 20 Segment position along the leaf  0 2 4 6 8 10 12 14 16 18 20 Segment position along the leaf  Fig. 3.4 Distribution of individual wax compound classes along the second leaf of rye at four growth stages. Coverages of (A) primary alcohols, (B) alkyl esters, (C) aldehydes, (D) alkanes, (E) secondary alcohols and (F) alkylresorcinols within the wax mixtures are given as averages of five independent parallels with standard deviations.  45  higher concentrations at growth stage IV than III. At growth stage III, ARs were also found starting only at 2 cm beyond the POE (segment 8), reaching a maximum of 0.1 µg/cm2 at 4 cm (segment 10) and then staying level until the tip of the blade (segment 11 to 20).  80  Percentages [% of total wax]  70 60 Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  50 40 30 20  .  Not indentified  Alkylresorcinols  sec. Alcohols  Alkanes  Aldehydes  Alkyl esters  prim. Alcohols  0  Fatty acids  10  Fig. 3.5 Relative amounts of compound classes in rye leaf waxes. Percentages of compound classes within the wax mixture on section 14 at four growth stages are shown as averages from five parallels with standard deviations.  Compared with the bulk waxes, the spatial distribution of ARs along the leaf axis indicated an even broader biosynthesis zone. The major wax production zone was within 2 cm beyond the POE, secondary alcohols accumulated mainly in zone a little broader (approximately 3 cm) beyond the POE, and ARs within roughly 4 cm beyond the POE. 46  Besides markedly different spatial distribution of ARs from those of all other wax constituents, timing of AR biosynthesis is distinct as well. Unlike most wax constituents, which are already present at the POE in early growth stages, ARs were not detected in younger leaves (at growth stage I); only two out of seven replicates at growth stage II contained traces of ARs; and then ARs constantly appeared at later growth stages (III and IV). Timing for most wax production was from each growth stage to its next adjacent stage. For ARs, it was between GS II (9-11 d) and GS IV (15-17 d).  47  3.3.4 Chain length distribution A total of 34 compounds were identified within the various compound classes, all of them containing fully saturated and unbranched alkyl chains. Homologous series of fatty acids, primary alcohols, aldehydes and alkyl esters were found to be largely dominated by compounds with even carbon numbers, with chain length ranges of C24 – C26, C24 – C28, C24-C26 and C40 – C46, respectively (Fig. 3.6). In contrast, alkanes with chain lengths C27 – C33, with a strong prevalence of odd-numbered homologs were detected, and secondary alcohols with C33chains. The secondary alcohol fraction was dominated by isomers with C-14 and C-16 hydroxylation. Overall, these patterns resemble those typically reported for other plant species, and confirm previous reports for rye leaf waxes. Homolog patterns for all the major compound  19 21 23 25  Fatty prim. Alkyl acids Alcohols esters  .  27 29 31 33  40 42 44 46  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  24 26 28  95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 20 15 10 5 0  24 26  Percentages [% of compound class]  classes stayed the same as Chapter 2 of rye leaf waxes.  Alkanes  Alkylresorcinols  Chain length Fig. 3.6 Chain length distributions within compound classes in rye leaf waxes. Percentages of individual homologs within each compound class are given for waxes on section 14 at four growth stages as averages from five parallels with standard deviations.  The fatty acid, primary alcohol and aldehyde fractions were all dominated by the C26 homolog, and the alkyl esters also showed a prevalence of isomers containing the C26 alcohol linked to C16 and C18 fatty acids. The alkanes had a relatively broad homolog distribution,  48  with a slight predominance of the C27 chain length. The relative amounts of the various homologs within these series of wax compounds did not vary significantly as a function of leaf development, for example comparing the wax compositions on segment 14 for the four growth stages (Fig. 3.6). Similarly, the chain length distributions did also not vary over the length of the leaf blade, when comparing the various segments at each developmental stage (data not  45 40 35 30 25 20 15 10 5 0  AR 19 AR 21 AR 23 AR 25  A  6  Percentages [%]  Percentages [%]  shown).  8  10  12  14  16  18  20  Segment position along the leaf  45 40 35 30 25 20 15 10 5 0  B  0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Fig. 3.7 Percentage of alkylresorcinol homologs within the alkylresorcinol fractions. Averages standard deviations from five independent parallels are given at growth stages III (A) and IV (B).  Within the alkylresorcinols, four different compounds were identified, defining a homologous series with side chains ranging from C19 to C25, with C23 as the maximum. Homologs with even-numbered side chains were detected, albeit only at trace levels. This chain length profile did not vary over time (Fig. 3.6) or as a function of the position along the leaf axis (Fig. 3.7).  49  3.4 Conclusions From the systematic time course analysis of wax composition of Secale cereale, the following conclusions can be drawn: (1) As for spatial distribution and wax accumulation zones: None of the wax components except alkanes were evenly distributed along rye leaves from POE to the tips of rye leaves. Primary alcohols, secondary alcohols, aldehydes and alkyl esters started low at POE followed by a rise over 2 - 3 cm until reaching a plateau. The wax accumulation zone could be inferred to be 2 - 3 cm beyond the POE for those compounds. ARs at growth stage III started with zero at the first two centimetres and then peaked at segment 4 to 10, followed by a drop to a plateau; at growth stage IV, ARs started to increase at the fourth centimetre after the POE, followed by a drop to a plateau. ARs had a broader production zone than other wax compounds, within 4 cm above the POE. (2) As for timing for wax production: Timing for most wax constituents was from each growth stage to its next adjacent stage. For ARs, it was between GS II (9 - 11 d) to GS IV (15 17 d). ARs were not formed at the same time as most other wax compounds.  50  References (1)  Richardson, A.; Franke, R.; Kerstiens, G.; Jarvis, M.; Schreiber, L.; Fricke, W.  Planta 2005, 222, 472-483. (2)  Bird, S. M.; Gray, J. E. New Phytologist 2003, 157, 9-23.  (3)  Kunst, L.; Samuels, A. L. Progress in Lipid Research 2003, 42, 51-80.  (4)  Rhee, Y.; Hlousek-Radojcic, A.; Ponsamuel, J.; Liu, D. H.; Post-Beittenmiller,  D. Plant Physiol. 1998, 116, 901-911. (5)  Richardson, A.; Wojciechowski, T.; Franke, R.; Schreiber, L.; Kerstiens, G.;  Jarvis, M.; Fricke, W. Planta 2007, 225, 1471-1481. (6)  Kavanova, M.; Lattanzi, F. A.; Grimoldi, A. A.; Schnyder, H. Plant Physiol.  2006, 141, 766-775.  51  Chapter 4: Conclusions and future work  This project, while focused on alkylresorcinols in Secale cereale, had further interests in larger questions of plant cuticular wax and alkylresorcinol biosynthesis. We are interested in the identification and characterization of an alkylresorcinol synthase, as well as discovering the role alkylresorcinols play in wax. Investigating how different wax components are arranged into epicuticular and/or intracuticular layers as well as monitoring when and where alkylresorcinols deposited in the leaf cuticle is another aspect of interest. Below is a review of some links witnessed among various parts of the research, possible insights into the questions raised and possible future experiments. From selective and non-selective analysis of cuticular wax in Secale cereale, it was concluded that (1) with regard to localization of ARs, 3% of ARs (with alkyl side chain lengths ranging from C19 to C27) in total waxes were discovered in the leaf cuticular wax of rye; ARs were limited mainly to the leaf cuticle, with quite low concentration found in the internal tissue; ARs were only found in the intracuticular wax layer while not detected in the epicuticular wax layer, (2) with regard to typical wax components, VLCFA derivatives, the total wax mixture is composed of primary alcohols (71%), alkyl esters (11%), aldehydes (5%), and small amounts of alkanes, steroids, secondary alcohols, fatty acids and unknown compounds. Abaxial and adaxial waxes contained very similar relative quantities of the same compound classes, and similar homologous constituents, making rye leaf an example where two surfaces of the same organ can share the same overall makeup-cuticular wax, even though the two surfaces demonstrate physiologically and ecologically different significance. From a systematic time course study on wax composition of Secale cereale, three major conclusions were drawn: (1) with respect to the spatial distribution, none of the wax components except alkanes were evenly distributed along rye leaves from POE to the end of the leaves. Alcohols and alkyl esters started low followed by a rise until reaching a plateau. ARs at growth stage III started with zero at the first two centimeters and then rose gradually to a plateau; at growth stage IV, ARs started to increase at the fourth centimeter after POE followed by a drop to a plateau. (2) With respect to the location of the growth/production zone: the growth zone above POE varied among different compound classes. The major wax production zone was within 2 cm above POE, secondary alcohol lied a little further and ARs was within 4 52  cm above POE. Based on this information, the location of highest ARS gene expression is inferred to be within 4 cm above POE. (3) With respect to the timing of wax production, most wax constituents were formed from each growth stage to its next adjacent stage. For ARs, accumulation rates peaked between GS II (9-11 d) to GS IV (15-17 d). ARs were not formed at the same time as most waxes. The chemical data obtained in this project provides important information to better design experiments towards ARS gene cloning. According to the results related to location and timing for AR production, the leaf segments at certain location (3-5 cm beyond POE of leaf two) and at the certain time (day 9-17 after germination of the plant) could be harvested for mRNA extraction, followed by cDNA synthesis and RT-PCR reactions leading eventually to the whole sequence of ARS genes. During the course of identification of ARs in wax, I found another homologous series of evenly spaced compounds similar to ARs. Their molecular ions also differed by 28 mass units, suggesting a homologous series differing by –CH2CH2- units. All of them showed peaks at m/z 73, 282 and 296. Both m/z 282 and 295 were 14 more than the characteristic peaks of alkylresorcinol TMSi ethers (268 and 281). Their corresponding fragments [M-15]+ indicated the loss of a methyl group from the TMSi derivatives. The molecular ions were 14 higher than the ARs eluting shortly before them, indicating a difference of one -CH2- unit from the ARs. The homologous series were inferred to consist of methyl-substituted ARs, yet the substitution position is not known. It could be ring-substituted, or alkyl chain-substituted at the first carbon next to aromatic ring. Future experiments are to identify those compounds by (a) synthesizing the potential candidates and then comparing their GC-MS behavior with the unidentified wax components, or (b) isolating and purifying the compounds for structure clarification using modern instrumental tools such as NMR, MS and IR. SEM has been widely used in cuticle studies, yet it was not involved in the current project. However, it would be interesting to see the surface changes on cuticle using SEM technique, especially in the major wax production zone (2 cm above POE) at relevant times (GS I (7-8 d) to GS II (9-11 d)). The larger question behind this project is about the function of alkylresorcinols in the cuticle. Since the majority of ARs are not located at the very surface of rye leaves, it is impossible that they could play a role in the direct interaction with insects or microorganisms. 53  However, they may function in regulating cuticle permeance. Whether alkylresorcinols moderate the permeance of the cuticle could be assessed by either of two ways: (1) heterologous expression of an ARS gene in an apropriate host plant; (2) analysis of permeance of cuticles of rye leaves at different ages. The effect of ARs on permeance could be evaluated by linking the amounts of ARs at any corresponding age to the water barrier effectiveness. In the same way, other wax constituents could be assessed for usefulness in preventing water loss. Sequencing and characterizing ARS genes continue in our lab. The ultimate goal of the project is to learn more about how ARS enzyme works and which amino acids or protein folding style determines substrate specificity, by comparing ARS with STS and CHS. Once ARS genes are successfully cloned, they can be characterized by yeast transformation, heterologous expression through Arabidopsis thaliana and over-expression of the gene in rye itself. The temporal and spatial expression patterns can be revealed by Northern blotting and in situ hybridization, respectively.1-3 Seeking out key amino acids by site directed mutagenesis might be a way to determine vital amino acids playing biosynthetic roles in the enzyme. Crystallization of the enzyme and X-ray crystallographic determination of its three-dimensional structure may explain substrate specificity of the ARS enzyme. The research work presented here has reached all its goals and provided detailed chemical insights into the accumulation of ARs in the cuticular waxes of rye leaves. This important information can contribute towards a better understanding of wax and AR biosynthesis. This is the first time that ARs as a wax family as well as its time course have been reported. It is expected that future research into the biological functions of cuticular ARs will build on the results presented here.  54  References (1)  Millar, A. A.; Kunst, L. Plant Journal 1997, 12, 121-131.  (2)  Popelka, J. C.; Altpeter, F. Molecular Breeding 2003, 11, 203-211.  (3)  Samach, A.; Kohalmi, S. E.; Motte, P.; Datla, R.; Haughn, G. W. Plant Cell  1997, 9, 559-570.  55  2.2 2.0 1.8 1.6 1.4 1.2 1.0 0.8 0.6 0.4  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  A  82  Percentage [%]  Percentage [%]  Appendix A  B  80 78 76 74 72 70 68  0  2  4  6  8 10 12 14 16 18 20  0  Percentage [%]  Percentage [%]  14  C  4 3 2 1 0  6  8 10 12 14 16 18 20  D  12 10 8 6 4  0  2  4  6  8 10 12 14 16 18 20  0  Segment position along the leaf 1.4  E Percentage [%]  5 4 3 2 1  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  6  Percentage [%]  4  Segment position along the leaf  Segment position along the leaf 5  2  F  1.2 1.0 0.8 0.6 0.4 0.2  0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Fig. A.1 Percentages of compound classes in total wax at various growth stages. (A) Fatty acids, (B) primary alcohols, (C) aldehydes, (D) alkyl esters, (E) alkanes and (F) secondary alcohols  56  0.7  A  B  0.6  3.5  0.5  3.0  0.4  2.5  0.3  2.0  0.2 0.1  1.5 0  2  4  6  0  8 10 12 14 16 18 20  Segment position along the leaf  2  96  Percentage [%]  C  96 95 95 94 94  4  6  8 10 12 14 16 18 20  Segment position along the leaf 1.0  97  Percentage [%]  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  Percentage [%]  Percentage [%]  4.0  D  0.8 0.6 0.4 0.2  93 0.0  93 0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Percentage [%]  1.9  E  1.8 1.7 1.6 1.5 1.4 1.3 1.2 0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Fig. A.2 Percentage of primary alcohol homologs in total primary alcohols. (A) tetracosanol, (B) pentacosanol, (C) hexacosanol, (D) heptacosanol, (E) octacosanol.  57  45  A  45  Percentage [%]  Percentage [%]  50  40 35 30 25 20  B  40 35 30 25 20 15 10  0  2  4  6  8 10 12 14 16 18 20  0  35  C  Percentage [%]  Percentage [%]  35 30 25 20 15  4  6  8 10 12 14 16 18 20  D  30 25 20 15 10  10  5 0  2  4  6  8 10 12 14 16 18 20  0  Segment position along the leaf 60  4  6  8 10 12 14 16 18 20  Segment position along the leaf 90  E  2  F  80  Percentage [%]  Percentage [%]  2  Segment position along the leaf  Segment position along the leaf  50  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  40  70  30  60  20  50  10 0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  40 0  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Fig. A.3 Percentage of individual alkane homologs and fatty acid homologs in total alkanes and total fatty acids, respectively. (A) heptacosane, (B) nonacosane, (C) hentriacontane, (D) tritriacontane, (E) tetracosanoic acid, (F) cerotic acid (26C).  58  Percentage [%]  A  7  Growth Stage I Growth Stage II Growth Stage III Growth Stage IV  6 5 4 3  Percentage [%]  28  8  2  B  26 24 22 20 18 16 14  0  2  4  6  8 10 12 14 16 18 20  0  Percentage [%]  Percentage [%]  C  0  2  4  6  8 10 12 14 16 18 20  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Segment position along the leaf 42 40 38 36 34 32 30 28 26 24 22 20  2  50 48 46 44 42 40 38 36 34 32  D  0  Segment position along the leaf  2  4  6  8 10 12 14 16 18 20  Segment position along the leaf  Fig. A.4 Percentages of individual ester homologs in total esters. (A) C40 alkylester, (B) C42 alkylester, (C) C44 alkylester, (D) C46 alkylester.  59  Table A.1 Statistical comparison of the accumulation of total wax at four growth stages. Listed data are P values at N=6 by one-way ANOVA.  Segment 14  GS I-14  GS II-14 GS III-14 GS IV-14 0.001864 0.009328 0.001841 0.8741 1 0.8715  GS I-15  GS II-15 GS III-15 GS IV-15 0.01609 0.2541 0.07341 0.4918 0.8805 0.8955  GS I-16  GS II-16 GS III-16 GS IV-16 0.3 0.9533 0.7223 0.5791 0.8709 0.9502  GS I-17  GS II-17 GS III-17 GS IV-17-18 0.1233 0.597 0.1498 0.7047 0.9996 0.7664  GS I-18  GS II-18 GS III-18 GS IV-17-18 0.319 0.9975 0.2791 0.4144 0.9997 0.3674  GS I-19  GS II-19 GS III-19 GS IV-20 0.9984 0.9796 0.9632 0.9441 0.9893 0.8224  GS I-20  GS II-20 GS III-20 GS IV-20 0.5623 0.898 0.4456 0.9227 0.9971 0.8416  GS I-14 GS II-14 GS III-14 GS IV-14 Segment 15 GS I-15 GS II-15 GS III-15 GS IV-15 Segment 16 GS I-16 GS II-16 GS III-16 GS IV-16 Segment 17 GS I-17 GS II-17 GS III-17 GS IV-17-18 Segment 18 GS I-18 GS II-18 GS III-18 GS IV-17-18 Segment 19 GS I-19 GS II-19 GS III-19 GS IV-20 Segment 20 GS I-20 GS II-20 GS III-20 GS IV-20  60  Table A.2 Statistical comparison of the accumulation of primary alcohols at four growth stages. Listed data are P values at N=6 by one-way ANOVA.  Segment 14  GS I-14  GS II-14 GS III-14 GS IV-14 0.00389 0.0117 0.002471 0.9556 0.9965 0.8864  GS I-15  GS II-15 GS III-15 GS IV-15 0.0493 0.5 0.1081 0.7563 1 0.7568  GS I-16  GS II-16 GS III-16 GS IV-16 0.3731 0.9922 0.6319 0.5273 0.9681 0.7915  GS I-17  GS II-17 GS III-17 GS IV-17-18 0.2837 0.7433 0.1102 0.8387 0.9431 0.5202  GS I-18  GS II-18 GS III-18 GS IV-17-18 0.2976 0.9862 0.1212 0.4701 0.9472 0.2169  GS I-19  GS II-19 GS III-19 GS IV-20 0.8846 0.975 0.876 0.9891 0.4735 0.6567  GS I-20  GS II-20 GS III-20 GS IV-20 0.7327 0.9436 0.3556 0.9633 0.9105 0.6722  GS I-14 GS II-14 GS III-14 GS IV-14 Segment 15 GS I-15 GS II-15 GS III-15 GS IV-15 Segment 16 GS I-16 GS II-16 GS III-16 GS IV-16 Segment 17 GS I-17 GS II-17 GS III-17 GS IV-17-18 Segment 18 GS I-18 GS II-18 GS III-18 GS IV-17-18 Segment 19 GS I-19 GS II-19 GS III-19 GS IV-20 Segment 20 GS I-20 GS II-20 GS III-20 GS IV-20  61  Table A.3 Statistical comparison of the accumulation of secondary alcohols at four growth stages. Listed data are P values at N=6 by one-way ANOVA. Segment 14  GS I-14  GS II-14 GS III-14 GS IV-14 0.002754 0.000318 0.000211 0.5817 0.2622 0.9277  GS I-15  GS II-15 GS III-15 GS IV-15 0.01163 0.004519 0.0003524 0.9711 0.2794 0.5043  GS I-16  GS II-16 GS III-16 GS IV-16 0.2078 0.09876 0.01474 0.9736 0.5305 0.7791  GS I-17  GS II-17 GS III-17 GS IV-17-18 0.06808 0.02342 0.002031 0.9526 0.3684 0.6688  GS I-18  GS II-18 GS III-18 GS IV-17-18 0.3748 0.1618 0.01251 0.9476 0.2856 0.5726  GS I-19  GS II-19 GS III-19 GS IV-20 0.9978 0.9156 0.4074 0.9662 0.5098 0.7807  GS I-20  GS II-20 GS III-20 GS IV-20 0.9089 0.5933 0.06148 0.93 0.2075 0.4907  GS I-14 GS II-14 GS III-14 GS IV-14 Segment 15 GS I-15 GS II-15 GS III-15 GS IV-15 Segment 16 GS I-16 GS II-16 GS III-16 GS IV-16 Segment 17 GS I-17 GS II-17 GS III-17 GS IV-17-18 Segment 18 GS I-18 GS II-18 GS III-18 GS IV-17-18 Segment 19 GS I-19 GS II-19 GS III-19 GS IV-20 Segment 20 GS I-20 GS II-20 GS III-20 GS IV-20  62  Table A.4 Statistical comparison of the accumulation of alkyl esters at four growth stages. Listed data are P values at N=6 by one-way ANOVA.  Segment 14  GS I-14  GS II-14 GS III-14 GS IV-14 0.0182 0.04369 0.01384 0.9745 0.9993 0.9461  GS I-15  GS II-15 GS III-15 GS IV-15 0.03274 0.1143 0.1168 0.9195 0.9154 1  GS I-16  GS II-16 GS III-16 GS IV-16 0.09086 0.6701 0.4496 0.5288 0.7497 0.9816  GS I-17  GS II-17 GS III-17 GS IV-17-18 0.3819 0.875 0.7924 0.8113 0.8901 0.9982  GS I-18  GS II-18 GS III-18 GS IV-17-18 0.8729 0.9922 0.9413 0.9633 0.9975 0.9917  GS I-19  GS II-19 GS III-19 GS IV-20 0.9538 1 0.7697 0.948 0.9689 0.7573  GS I-20  GS II-20 GS III-20 GS IV-20 0.8197 0.5359 0.4312 0.9594 0.9043 0.9978  GS I-14 GS II-14 GS III-14 GS IV-14 Segment 15 GS I-15 GS II-15 GS III-15 GS IV-15 Segment 16 GS I-16 GS II-16 GS III-16 GS IV-16 Segment 17 GS I-17 GS II-17 GS III-17 GS IV-17-18 Segment 18 GS I-18 GS II-18 GS III-18 GS IV-17-18 Segment 19 GS I-19 GS II-19 GS III-19 GS IV-20 Segment 20 GS I-20 GS II-20 GS III-20 GS IV-20  63  Table A.5 Statistical comparison of the average wax loads of alkanes at four growth stages. Listed data are P values at N=6 by one-way ANOVA.  GS I GS I GS II GS III GS IV  GS II GS III GS IV 0.9306 0.000217 0.000166 0.000172 0.000166 0.000577  64  Appendix B: Cloning of alkylresorcinol synthase (ARS) genes  B.1: Summary of cloning procedures B.1.1 Strategies for ARS gene cloning (1) Primers designed from STS/CHS sequences Protein sequences of CHS and STS enzymes originating from various plants were aligned. Conserved regions of protein sequences of STS and CHS enzymes were located, respectively. Some of STS consensus was also well conserved in CHS, while some were not. Those STS well-conserved regions yet not conserved in CHS were considered to be the characteristic regions of STS, and therefore were chosen for primer designing. Four pairs of primers were designed based on the STS from peanut (Arachis hypogaea), grape (Vitis vinifera) and Arabidopsis (Arabidopsis thaliana) (Table B.1) and one pair from rice (Oryza sativa) (Table B.2).  Table B.1 Primers designed based on the consensus of STS/CHS from peanut (Arachis hypogaea), grape (Vitis vinifera) and Arabidopsis (Arabidopsis thaliana).  Primer Name F1 F2 F3 F4 R1  Primer Sequence GTVTSTAYCAGTCWGAYTWYGC GTRTYGATCAGAGYACATATGC MWARCVWDKGYGCWTAYAWGGC CAAACATTGGTGCTTATATGGC ATRSWWATACCMARTGGRTC  Notes  From Conserved region of STS which are not conserved in CHS  Table B.2 Primer designed based on the consensus of STS/CHS from rice (Oryza sativa). Primer Name osPKS-F  Primer Sequence ATGGCACCTGTTCCGGCCAC  osPKS-R  TTAATTTCCCTTGAGGCCGCTTC  Notes From 34 STS/CHS sequences in rice (Oryza sativa)  65  (2) Primers designed from ARS ARS gene discovered in bacteria (Azotobacter vinelandii) is not used to design primers due to its low identity (20 a. a. %) with CHS or STS genes.1 Colleagues from USDA kindly provided three ARS genes from rice (Oryza sativa) and two from sorghum (Sorghum bicolor). The two sorghum ARS match well (70% a. a. identity) with rice ARS sequences. Using the five sequences at hand, four pairs of primers were designed (Table B.3).  Table B.3 Primer designed based on the consensus of three ARS from rice (Oryza sativa) and two from sorghum (Sorghum bicolor).  Primer Name ARS-F ARS-R ARS F1 ARS F2 ARS F3 ARS R1  Primer Sequence ATCGCCATCGGCACTGCAAACCC GCACCACTCATGTTCCCAAACTC CCACGGTSCTMGCMATCGGCAC CGACTAYVCSGACTACTACTTC AAGGCGATCRMBGAGTGGGGCC ACGCGYGCRCCGCGGTTGTTCTC  Notes Designed from 3 rice ARS, 2 sorghum ARS  The two sorghum ARS sequences were used to blast similar segments in rye (Secale cereale) gene database. Based on the searching results, three pairs of primers were designed to arrest ARS in ryes (Table B.4).  Table B.4 Primer designed based on the consensus of blasted sequences using the two sorghum (Sorghum bicolor) ARS sequences to rye sequence database.  Primer Name Rye1-F Rye1-R Rye2-F Rye2-R Rye3-F Rye3-R  Primer Sequence TCAAGATCACCAAGAGCGACCACA TCAGGTTTACCTTTGCCTCGACCA TCAGGTTTACCTTTGCCTCGACCA TAGCCAAAGACCTCGCTGAGAACA AGGACAACTGTCTCCACAGTGA CTCATCTCCAAGAACATCGAGC  Notes From 2 sorghum ARS blasting in rye database  66  B.1.2 Procedures of ARS gene cloning (1) mRNA extraction Firstly, plant materials were homogenized and mRNA was extracted. 100 mg of rye leaves was grounded in 1 mL DNAlater and then 1000 µL TRI Reagent. The paste was left at room temperature for 5 min before centrifuging at the speed of 11,000 rpm at 4 °C for 10 min. The supernatant was transferred into a clean RNase-free tube. Secondly, the RNA extracts were pre-cleaned by adding 100 µL of CHCl3, vortex mixing, incubating at room temperature for 15 min followed by centrifuging at about 11,000 rpm at 4 °C for 15 min. The aqueous phase was transferred into a fresh RNase-free tube. Thirdly, mRNA was precipitated and cleaned. 500 µL of isopropanol was added and vortex mixed for 10 s and then incubated at room temperature for 10 min. The mixture formed was centrifuged at 11,000 rpm at room temperature for 8 min. Supernatant was discarded and 1 mL of 75% of ethanol in water was added to the precipitates. After centrifugation at 6000 rpm at room temperature for 5 min, ethanol was removed with pipette and the leftover was air-dried for less than 10 min. Finally, the concentrated and cleaned mRNA extracts was dissolved in 30 µL of deionized and autoclaved water and stored in -20 °C freezer.  (2) Reverse transcription from mRNA to cDNA 10 µL of mRNA mixed with 1 µL of Oligo (dt)23 was incubated at 70 °C for 5 min followed by chilling on ice. Then 2 µL of reaction buffer, 0.5 µL of RNase inhibitor and 3.5 µL of nuclease-free water were added sequentially. The resulting mixture was incubated at 37 °C for 5 min before 1 µL of reverse transcriptase (200 ng/µL) was added to the mixture to start cDNA synthetic reaction. The reaction proceeded at 42 °C for 60 min before it was stopped by heating in 70 °C water batch for 10 min. The quenched reaction mixture was chilled on ice before storing at -20 °C.  (3) PCR reactions Using the primers listed in the tables, pcr was performed with taq polymerase (NEB) using 2 µL template, 1 µL of each primer, 2 µL of reaction buffer, 0.4 µL of dNTP, 13.4 µL of water (deionized and autoclaved) with the program: 94 °C- 3min, 10x (94 °C-15s, 65°C-30s, 72°C-20s), 20x (94°C-15s, 55°C-30s, 72°C-45s), 72°C-7min. pcr reaction was performed on a 67  Mastercycler Gradient thermocycler (Eppendorf, Germany). Subsequent analysis was performed with gel electrophoresis (1% LiCH3COO, 0.7% agarose, 200V, ~15 min, DNA visualized using SYBR® Gold Nucleic Acid Gel Stain under UV illumination).  B.2: Problems encountered Although I managed several times to design various primers, RT-PCR did not give the right products. Primers in Table B.1 were designed from a conserved region of STS yet not conserved in CHS. Those STS and CHS were picked from plants such as peanut (Arachis hypogaea), grape (Vitis vinifera) and Arabidopsis (Arabidopsis thaliana), which may not have enough similarity in CHS and/or STS sequences with rye (Secale cereale). Hence, the failure of those primers may be inferred to be due to a lack of similarity of the STS sequences with those in rye. Therefore primers in Table B.2 were designed according to the STS/CHS sequences sourced from rice (Oryza sativa). However, repeated efforts to synthesize those targeted pcr products were unsuccessful. At the same time, chemical analysis showed that the very tender leaves at Growth Stage I did not contain ARs, indicating that ARS enzyme was inactive at that time point. It might explain why no PCR products were found from the leaves at Growth Stages I. Thus, tender leaves at Growth Stage II, III and IV were used instead. The primers in Table B.1 and B.2 were re-used for harvesting pcr products, unfortunately without positive results. Colleagues at USDA gave me hope by kindly providing five ARS genes (three from rice and two from sorghum). Aligning the five sequences, I designed four pairs of primers according to the consensus. In addition, another three pairs of primers were designed by blasting the two sorghum ARS sequences in rye gene database. Unfortunately, none of those primers worked after repeated efforts.  68  Reference (1)  Funa, N.; Ozawa, H.; Hirata, A.; Horinouchi, S. Proc. Natl. Acad. Sci. U. S. A.  2006, 103, 6356-6361.  69  

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