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Synthesis of very-long-chain bifunctional and isotope-labeled compounds for biochemical investigations… Peng, Chen 2012

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 SYNTHESIS OF VERY-LONG-CHAIN BIFUNCTIONAL AND ISOTOPE-LABELED COMPOUNDS FOR BIOCHEMICAL INVESTIGATIONS INTO NOVEL COMPOUNDS IN PLANT CUTICULAR WAXES   by  Chen Peng  B.Sc.(Honors), Peking University, 2009  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)   December 2012 © Chen Peng, 2012 ii  Abstract Plant cuticles are the interface for plant-environment interactions, and the first barrier protecting plants from environmental stresses such as water loss and pathogens. Structurally, the cuticle consists of a hydrophobic polymer lattice, cutin, and cuticular waxes deposited inside and outside cutin. The major components of the cuticular waxes are aliphatics derived from very-long-chain (VLC) fatty acids, such as alkanes and aldehydes. Besides compounds with primary functionalities, some compounds with two or more functional groups have also been identified in cuticular waxes. However, only limited knowledge about them has been acquired so far.  The current work was to identify novel 1,2- and 1,3-bifunctional wax compounds from various plant species, in order to expand our current understanding of their structure, biosynthesis and function in plant cuticles. Synthetic methods were first developed to produce various VLC 1,2- and 1,3-bifunctionalized standard compounds for current and future structure elucidation studies. Unknown compounds found in Cosmos bipinnatus petal wax were identified as alkane-1,2-diol monoacetates by GC-MS, with chain lengths ranging from C20 to C24. The ratio between the primary and the secondary monoacetates was quantified to be 3:5, as opposed to the thermodynamic equilibrium ratio of 7:3. Novel β-hydroxy acid methyl esters were also identified from Aloe arborescens leaf wax, with chain lengths ranging from C26 to C30. In addition, two NMR-based methods were established to study the stereoconfigurations of alkane-1,2-diols from C. bipinnatus petal wax, and the carbons bearing secondary hydroxyl functionality were determined to have predominately the R-configuration.  Apart from the research on bifunctional compounds, synthetic methods to produce β-deuterium labeled VLC substrates were also established. The resulting C30 fatty acid methyl ester was double-labeled and can be used directly or indirectly as substrates in future biochemical assays. At last, the hypothetic substrate for the CER1 enzyme implicated in wax alkane biosynthesis, C30 aldehyde, was synthesized and used in in vivo assays with heterologously expressed protein. iii  Preface A version of Chapter 3 has been published as: Buschhaus, C., Peng, C., Jetter, R., Very-long-chain 1,2- and 1,3-bifunctional compounds from the cuticular wax of Cosmos bipinnatus petals. Phytochemistry (2012), http://dx.doi.org/10.1016/j.phytochem.2012.07.018  The work in Chapter 3 was based on previous research performed by C. Buschhaus, who prepared the wax sample, obtained and analyzed the mass spectra of the unknown compounds and proposed possible structures based on the mass spectra.  The work described in Chapter 5 was conducted partially in collaboration with A. Luna and R. Racovita. The wax material was provided by A. Luna, and the mass spectra of the unknown compounds were obtained by A. Luna and R. Racovita.  The yeast feeding experiments in Chapter 6 were performed in collaboration with M. Skvortsova. She conducted the gene transformation, PCR experiments and yeast cell cultivation. I was responsible for the synthesis of C30 aldehyde substrate. We both made equal contributions to the yeast feeding, the lipid extraction and the following sample analysis.  All the remaining syntheses, experiments and data analyses were my contribution. I also wrote the manuscript of this thesis. Thesis revision was performed with the help of Dr. R. Jetter.      iv  Table of content Abstract…………………………………………………………………………………………...ii Preface…………………………………………………………………………………………...iii Table of content ............................................................................................................. iv Acknowledgement ........................................................................................................ vii List of Tables ............................................................................................................... viii List of Figures ................................................................................................................ ix List of Schemes ............................................................................................................ xii List of Abbreviations .................................................................................................... xiii Dedication .................................................................................................................... xv 1   Introduction to plant cuticles and biosynthesis of wax compounds .......................... 1 1.1 Composition and structure of plant cuticles ..................................................... 1 1.2 Biosynthesis of very-long-chain fatty acid ........................................................ 2 1.3 The biosynthesis of mono-functionalized wax compounds............................... 4 1.3.1 The primary alcohol pathway .......................................................................... 5 1.3.2 Biosynthesis of alkanes................................................................................... 6 1.4 Biosynthesis of wax compounds with multiple functional groups ..................... 8 1.4.1 Symmetric secondary alcohols, ketones, ketols and diols ............................. 10 1.4.2 β-Diketones and hydroxy-β-diketones ........................................................... 12 1.4.3 Primary/secondary bifunctional wax compounds ........................................... 13 1.4.4 Asymmetric secondary alcohols, ketones and secondary/secondary diols…………………………………………………………………………………..14 1.5 Objectives ..................................................................................................... 15 2   Materials and methods ......................................................................................... 17 2.1 Plant materials and wax extraction ................................................................ 17 2.2 Sample derivatization .................................................................................... 17 2.3 GC-MS/FID analysis ...................................................................................... 17 2.4 Chemical synthesis........................................................................................ 18 2.5 Yeast feeding experiment and lipid extraction ................................................ 18 2.5.1 Yeast expression and feeding conditions ...................................................... 18 2.5.2 Yeast total lipid extraction.............................................................................. 19 3   Identification and quantification of 1,2-diol monoacetates in Cosmos bipinnatus petal wax ...................................................................................................................... 20 3.1 Introduction ................................................................................................... 20 3.2 Results .......................................................................................................... 23 3.2.1 Synthesis of alkane 1-hydroxy-alkane-2-one standards ................................ 23 v  3.2.2 Synthesis of 1,2-diol monoacetates standards .............................................. 25 3.2.3 Study of the acyl-migration behavior of the Cosmos bipinnatus 1,2-diol monoacetates ............................................................................................... 33 3.2.4 Analysis and quantification of 1,2-diol monoacetates in C. bipinnatus petal wax...…………………………………………………………………………………34 3.3 Discussion ..................................................................................................... 36 4   NMR-based determination of the stereoconfigurations of cosmos 1,2-Diols ......... 39 4.1 Introduction ................................................................................................... 39 4.2 Results .......................................................................................................... 41 4.2.1 Synthesis of S- and R-1,2-docosanediol standards and isolation of 1,2-diols from C. bipinnatus petal wax ......................................................................... 41 4.2.2 Stereochemical investigation of 1,2-bifunctional wax compounds using Mosher’s Method .......................................................................................... 43 4.2.3 Study of the stereochemistry of 1,2-bifunctional wax compounds using boronic acid-based method. ...................................................................................... 47 4.3 Discussion ..................................................................................................... 51 4.3.1 Comparison of the two derivatization methods .............................................. 51 4.3.2 Possible biosynthetic pathway to 1,2-diols in cosmos ................................... 52 4.3.3 Possible biosynthetic pathway to 1,2-diol monoacetates in cosmos .............. 55 5   β-Keto acid derivatives: potential pathway intermediates and end products ......... 57 5.1 Introduction ................................................................................................... 57 5.2 Results .......................................................................................................... 60 5.2.1 Synthesis of β-functionalized wax compounds .............................................. 60 5.2.2 Identification of β-functionalized wax compounds in Aloe arborescens leaf waxes………………………………...………………………………………………64 5.3 Discussion ..................................................................................................... 68 5.3.1 Synthesis of β-functionalized wax compounds .............................................. 68 5.3.2 Biosynthesis of β-hydroxy acid methyl esters in A. arborescens ................... 69 6   Synthesis of isotope-labeled very-long-chain aliphatic substrates and preliminary in vivo assays of CER1 in yeast (Saccharomyces cerevisiae) ................................. 72 6.1 Introduction ................................................................................................... 72 6.2 Results .......................................................................................................... 73 6.2.1 Synthesis of deuterium-labeled very-long-chain substrate ............................ 74 6.2.2 Synthesis of very-long-chain fatty aldehyde .................................................. 78 6.2.3 Preliminary in vivo assays of CER1 using heterologous expression in yeast………………………………………………………………………………….79 6.3 Discussion ..................................................................................................... 82 6.3.1 Synthesis of double-deuterium labeled VLC substrates ................................ 82 6.3.2 Preliminary in vivo assays of CER1 in yeast ................................................. 83 7   Summary and future directions ............................................................................. 85 vi  7.1 Synthesis of 1,2- and 1,3-bifunctional standard compounds .......................... 86 7.2 Identification of novel 1,2- and 1,3-bifunctional wax compounds.................... 87 7.3 Determination of the stereoconfigurations of cosmos 1,2-diols ...................... 87 7.4 Synthesis of isotope-labeled substrates ......................................................... 88 References .................................................................................................................. 90 Appendices ................................................................................................................ 106 Appendix A: Synthetic protocols of the compounds in Chapter 3 ............................ 106 Appendix B: Synthetic protocols of the compounds in Chapter 4 ............................ 111 Appendix C: Synthetic protocols of the compounds in Chapter 5 ............................ 113 Appendix D: Synthetic protocols of the compounds in Chapter 6 ........................... 122                      vii  Acknowledgement I take this opportunity to express my sincerest gratitude and deep regards to my research supervisor, Dr. Reinhard Jetter, for his immense guidance, instruction and support throughout the course of my study. Without his substantive help and contribution this project would not have materialized.  I would like to thank all my lab mates, including Dr. Christopher Buschhaus, Mariya Skvortsova, Luke Busta, Ruonan Yao, Yan Cao, Alvaro Luna, Radu Racovita and Daniela Hegebarth, for providing extremely helpful assistance and sharing constructive advice. Their friendships have made the Jetter lab an enjoyable work place.  Finally, my appreciation also goes to my cherished friends and beloved parents, for their constant encouragement and always being a source of inspiration and support.   viii  List of Tables Table 3.1 Chain-length specific ratios of secondary monoacetates and primary monoacetates. Results were averaged between two independent samples. ....................................... 36   ix  List of Figures Figure 1.1 Diagram of the transverse view of plant cuticles and epidermal cells. This diagram was modified after Jetter et al 2000 and Jeffree 1996. 6 ................................................. 2 Figure 1.2 Reactions and enzymes involved in the very-long-chain fatty acid elongation cycle.10 ........................................................................................................................... 3 Figure 1.3 Biosynthesis of mono-functionalized wax compounds and secondary alcohols and ketones. The VLC acyl-CoAs are the substrates for both the primary alcohol pathway and the alkane pathway. Primary alcohols and esters are produced in the primary alcohol pathway, whereas aldehydes, alkanes, secondary alcohols and ketones are produced in the alkane pathway. .................................................................................... 5 Figure 1.4 Examples of multifunctional wax compounds ....................................................... 9 Figure 1.5 The hypothetic pathway leading to symmetric diols and ketols. The symmetric diols and ketols can be biosynthesized from the symmetric secondary alcohols and ketones by a MAH1-like hydroxylase. .......................................................................... 12 Figure 1.6 The hypothetic pathway leading to hentriacontane-14,16-dione. The β-diketone motif can be introduced by omitting the activities of KCR, HCD and ECR in two consecutive elongation cycles. ..................................................................................... 13 Figure 3.1 Mass spectra of diols from C. bipinnatus petal waxes.51 (A) Bis-TMSi ether of 1,2-docosanediol from series A1; (B) Bis-TMSi ether of 1,3-docosanediol from series A2.  .................................................................................................................................... 21 Figure 3.2 Mass spectra of unknowns in series B1 and B2.51 (A) TMSi ether of the most abundant compound in series B1; (B) TMSi ether of the most abundant compound in series B2. ..................................................................................................................... 22 Figure 3.3 Mass spectra of 1-hydroxy-alkan-2-one standards. (A) TMSi ether of 1-hydroxyoctan-2-one; (B) TMSi ether of 1-hydroxytetradecan-2-one. ......................... 25 Figure 3.4 GC-MS chromatogram of the product mixture from the acetylation of tetradecane-1,2-diol. .................................................................................................... 26 Figure 3.5 Mass spectra of acetates synthesized from 1,2-tetradecanediol.  (A) TMSi ether of 1-hydroxytetradec-2-yl acetate; (B) TMSi ether of 2-hydroxytetradecyl acetate. ....... 27 Figure 3.6 Characteristic 1H-NMR signals of isolated 2-hydroxydocosyl acetate and 1-hydroxydocosan-2-yl acetate. ................................................................................... 29 Figure 3.7 Co-elution experiment of the TLC fraction that contains 1,2-diol monoacetates and 1,2-docosanediol monoacetate standards. The most abundant peak in series B1 had approximately the same retention time as 3-6, and similarly the most abundant peak in series B2 had almost the same retention time as 3-5. .................................................. 31 x  Figure 3.8 Mass spectra of two representatives each from series B1 and B2.  (A) Identified as TMSi ether of 1-hydroxyeicosan-2-yl acetate; (B) identified as TMSi ether of 2-hydroxyeicosyl acetate; (C) identified as TMSi ether of 1-hydroxytetracosan-2-yl acetate; (D) identified as TMSi ether of 2-hydroxytetracosyl acetate. For the mass spectra of TMSi derivatized 1-hydroxydocosan-2-yl acetate and 2-hydroxydocosyl acetate, refer to Figure 3.2. .......................................................................................... 32 Figure 3.9 Study of acyl migration using 1H-NMR. Acyl migration was observed from day 1 (D1) to day 9 (D9) for both primary monoacetate (3-5) and secondary monoacetate (3-6) in CDCl3 solution. ........................................................................................................ 34 Figure 3.10 GC-MS chromatogram of the TLC fraction containing 1,2-diol-monoacetates. Abbreviations: ac-acid, ol-alcohol, sm-secondary monoacetate, pm-primary monoacetate. ............................................................................................................... 35 Figure 3.11 Relative amounts of 1,2-diol monoacetates (% of C22 diol 2-acetate) extracted from C. bipinnatus petal. Results were averaged between two independent samples. . 36 Figure 4.1 GC-FID chromatogram of purified 1,2-diols from C. bipinnatus petal wax. Peak abundances were normalized according to C24 alkane internal standard (set as 100). . 43 Figure 4.2 Characterization of 1,2-tetradecanediol 1-S-MTPA-ester 4-3. (A) 1H-NMR spectrum (400 MHz) of 4-3; (B) Mass spectrum and (C) major fragmentations of the TMSi ether of 4-3. ........................................................................................................ 45 Figure 4.3 Signature 1H-NMR (400 MHz) signals of the 1,2-diol 1-S-MTPA esters. ............. 46 Figure 4.4 NMR spectrum (300 MHz) of 4-8. ....................................................................... 49 Figure 4.5 Signature 1H-NMR (400 MHz) signals of the 1,2-diols derivatized using boronic acid based three-reagent system. ................................................................................ 50 Figure 4.6 Hypothetical biosynthesis pathways to 1,2-diols and 1,2-diol-monoacetates. 1,2-Diols can be biosynthesized by epoxidation of alkenes followed by hydrolysis, or by α-hydroxylation of fatty acids followed by reduction of the acid group into hydroxyl group. 1,2-Diol-monoacetates can be biosynthesized by esterification directly from 1,2-diols, or by esterificaion of the α-hydroxy acid, followed by reduction. ....................................... 55 Figure 5.1 Hypothetic alkane and 1,3-alkanediol biosynthesis pathways via β-ketoacyl-CoA intermediates. The β-keto acid can be derived from the elongation cycle and undergo instantaneous decarboxylation to afford alkan-2-one. The alkan-2-one can be reduced to alkane by a mechanism similar to the elongase reduction sequence. The reduction of β-ketoacyl-CoA to 1,3-diol can be achived by two enzymes: CER4-like and KCR-like reductases. Alternatively, alkane and 1,3-alkanediol can also be biosynthesized via fatty acid acyl-CoA intermediates (colored in grey). ............................................................. 58 Figure 5.2 Mass spectra of the TMSi ethers of C30 standard compounds . .......................... 64 Figure 5.3 GC-MS chromotograms of the TMSi ethers of Aloe waxes. (A) Full scan mode, (B) xi  SIM mode monitoring fragment m/z 175. ...................................................................... 65 Figure 5.4 Mass spectra of unknown compounds from A. arborescens leaf waxes. (A) TMSi ether of the first unknown homologue. (B) TMSi ether of the second unknown homologue. (C) TMSi ether of the third unknown homologue. ......................................................... 66 Figure 5.5 Co-elution expermient of the A. arborescens leaf wax and β-hydroxy acid methyl ester standard 5-15. The retention times of the third unknown homologue and 5-15 were approximately the same on GC-FID. ............................................................................ 67 Figure 5.6 Hypothetical biosynthesis pathways to β-hydroxy acid methyl esters. The methyl ester can be synthesized by an methyltransferase or acyltransferase from the corresponding acyl-CoA. The β-functional group can be introduced from the elongation or be installed via hydroxylation. .................................................................................. 70 Figure 6.1 Charaterization of deuterium-labeled substrate 6-4. (A) 1H-NMR spectrum (300 MHz) of 6-4; (B) The molecular ions of 6-4 (red bars) and the corresponding unlabeled ester (blue bars) in their mass spectra. ........................................................................ 76 Figure 6.2 Charaterizations of double-deuterium labeled substrate 6-7. (A) NMR spectrum (300 MHz) of 6-7; (B) molecular ions of 6-7 (red bars) and the corresponding unlabeled ester (blue bars) in their mass spectra. ........................................................................ 78 Figure 6.3 Co-elution expermient of the preliminary in vivo assays of CER1 in yeast. GC-FID chromatograms of assays with (A) C30 aldeyde fed to yeast expressing CER1; (B) C30 primary alcohol fed to yeast expressing CER1; (C) yeast expressing CER1 without substrate feeding; (D) wild-type yeast alone. (E) C29 alkane standard. ......................... 81 Figure 6.4 The molecular ions of 6-5 (red bar) and 5-15 (blue bar) in their mass spectra. ... 83   xii  List of Schemes Scheme 3.1 Synthesis of 1-hydroxy-2-ketone standards. .................................................... 24 Scheme 3.2 Synthesis of 1, 2-diol-monoacetate standards. ................................................ 26 Scheme 3.3 Synthesis of 2-hydroxytetradecyl acetate from 3-4. ......................................... 28 Scheme 3.4 Fragmentation patterns of the TMSi ethers of 1,2-docosanediol monoacetates.  .................................................................................................................................... 30 Scheme 4.1 Chiral derivatization agents and reactions for Mosher’s method. ..................... 40 Scheme 4.2 Boronic acid-based three-reagent derivatization system.................................. 41 Scheme 4.3 Synthesis of S- and R-1,2-docosanediol standards. ........................................ 42 Scheme 4.4 Synthesis of mono-MTPA-derivatized 1,2-diols................................................ 43 Scheme 4.5 Derivatization of 1,2-diols with the boronic acid-based three-reagent system. . 48 Scheme 5.1 Synthesis of octadecanoyl-Meldrum’s acid 5-3 and its derivatives. .................. 61 Scheme 5.2 Synthesis of long-chain β-functionalized compounds from methyl 3-oxoeicosanoate 5-5. .................................................................................................. 62 Scheme 5.3 Synthesis of octacosanoyl-Meldrum’s acid 5-11 and methyl 3-oxotriacontanoate 5-12. ............................................................................................................................ 62 Scheme 5.4 Synthesis of very-long-chain β-functionalized compounds from methyl oxotriacontanoate 5-12. ............................................................................................... 63 Scheme 5.5 Fragmentation patterns of methyl 3-hydroxytriacontanoate. ............................ 67 Scheme 6.1 Retrosynthetic analysis of β-D-labeled substrate. ............................................ 74 Scheme 6.2 Isotope-labeling strategy via β-iodide intermediate. Only elimination product was obtained. ...................................................................................................................... 75 Scheme 6.3 Isotope-labeling strategy via β-tosyl intermediate. ........................................... 75 Scheme 6.4 Synthesis of double-deuterium labeled very-long-chain substrate. .................. 77 Scheme 6.5 Synthesis of long-chain and very-long-chain fatty aldehydes. .......................... 79   xiii  List of Abbreviations ACP acyl carrier protein AMA 2-(anthracen-9-yl)-2-methoxyacetic acid BSTFA bis-N,O-(trimethylsilyl)trifluoroacetamide δ chemical shift CER eceriferum CDA chiral derivatization agent CoA coenzyme A d doublet D deuterium DCC N,N'-dicyclohexylcarbodiimide DCM dichloromethane DMAP 4-dimethylaminopyridine DMSO dimethyl sulfoxide ee enantiomeric excess ECR enoyl-CoA reductase EDC 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide) ER endoplasmic reticulum FAE fatty-acid-elongase FAR fatty acyl reductase FID flame ionization detection G3P glycerol-3-phosphate GC gas chromatography GPAT glycerol phosphate acyltransferase HCD β-hydroxyacyl-CoA dehydratase J coupling constant KCR β-ketoacyl-CoA reductase KCS β-ketoacyl-CoA synthase xiv  LACS long-chain acyl-CoA synthetase LAH Lithium aluminum hydride m multiplet mol mole(s) MS mass spectrometry MTA methoxyphenyl acetic acid MTPA α-methoxy-α-trifluoromethylphenylacetic acid MTPA-Cl α-methoxy-α-trifluoromethylphenylacetic acid chloride NAC n-acetylcysteine NMR Nuclear magnetic resonance PCC pyridinium chlorochromate PCR polymerase chain reaction q quartet Rf retention factor rt room temperature s singlet SAM S-adenosyl methionine t triplet TEA triethylamine THF tetrahydrofuran TLC thin layer chromatography TMSi trimethylsilyl Ts p-toluenesulfonyl UV ultraviolet VLC very-long-chain VLCFA very-long-chain fatty acid WS/DGAT wax synthase/diacylglycerol acyl transferase  xv  Dedication  To the lovely summer in Vancouver 1  1 Introduction to plant cuticles and biosynthesis of wax compounds The cuticle is a hydrophobic layer that covers the surface of primary aerial organs of land plants. Produced by epidermal cells, the cuticle plays a pivotal role in protecting plants against biotic and abiotic stresses. Most importantly, the cuticle helps to prevent non-stomatal water loss,1 which is believed to be critical to the evolution of land plants. It also protects the plants against the attack of pathogenic microbes and insect herbivores, reduces dust and other particles deposited onto the plant surfaces, and alleviates high-dose ultraviolet radiation from the sun.2  In this chapter, the chemical compositions of plant cuticle will first be introduced (section 1.1), followed by a discussion of the biosynthetic pathways leading to the various constituents of cuticular waxes. In particular, the elongation cycle generating very-long-chain fatty acids (section 1.2), and the pathways modifying them into all other wax compounds (sections 1.3 and 1.4) will be discussed. More specifically, section 1.3 will focus on the biosynthesis of wax compounds with primary functionalities, and the biosynthesis of the bi- and multifunctional wax compounds will be described in section 1.4.  1.1 Composition and structure of plant cuticles In general, the composition of the cuticle is categorized into two major components, based on their solubilities in organic solvents. The insoluble component is called cutin (Figure 1.1), which is a polymeric matrix of ω- and mid-chain hydroxy and epoxy C16 and C18 fatty acids connected directly by ester bonds and also through glycerol bridges.3 Cutin serves as the backbone of the cuticle and makes up 40-80% the total weight. The other major components in the cuticle are cuticular waxes, which can be categorized into aliphatics, alicyclics and aromatics. The aliphatics consist of very-long-chain fatty acids (VLCFAs) and derivatives, such as alcohols, esters, aldehydes and ketones.4 Apart from the esters, the chain lengths of other wax compounds usually range from C20 to C34. The chain lengths of the dimeric esters are between C38 and C70. Alicyclics and aromatics, mostly triterpenoids 2  and phenolics,5 respectively, are usually occurring in low amounts.  Figure 1.1 Diagram of the transverse view of plant cuticles and epidermal cells. This diagram was modified after Jetter et al 2000 and Jeffree 1996. 6 The wax film exterior to cutin is called epicuticular wax, as opposed to intracuticular wax, which is buried inside the cutin lattice (Figure 1.1).6a Epicuticular wax can be mechanically peeled off using adhesives, whereas organic solvent extraction will remove both epi- and intracuticular waxes. Therefore, the compositions of epi- and intracuticular wax mixtures can be analyzed separately.7 The cyclic compounds, like triterpenoids and alkylresorcinols, together with VLC primary alcohols preferably reside in the intracuticular layer. In contrast, VLC fatty acids and alkanes tend to accumulate in the epicuticular layer.7 No big differences were observed for the chain length profiles of aliphatic compounds across the two wax layers. The origin of this partition of wax compounds is yet to be discovered.  1.2 Biosynthesis of very-long-chain fatty acid Very-long-chain fatty acids are part of the wax compounds, and they serve as the precursors of other wax compounds, such as primary alcohols, alkanes, secondary alcohols and esters. In Arabidopsis thaliana, they are biosynthesized via the elongation of long-chain fatty acyl-CoAs. 3   Before the long-chain fatty acids enter the elongation process, they need to be liberated from the acyl carrier proteins (ACP) by acyl-ACP thioesterase, and then get converted to acyl-CoAs by long-chain acyl-CoA synthetase (LACS).8 In the elongation process, C2 moieties are added to the acyl-CoA substrate sequentially. This process is catalyzed by enzyme complexes localized to the endoplasmic reticulum (ER), called fatty-acid-elongase (FAE). Each FAE is a heterotetramer, consisting of four distinct enzymes. Similar to the synthesis of long-chain fatty acids, each elongation cycle consists of four consecutive enzymatic reactions (Figure 1.2).9 Firstly, the fatty acyl-CoA substrate is condensed with a malonyl-CoA catalyzed by β-ketoacyl-CoA synthase (KCS). The reduction of the β-keto group will produce β-hydroxy acyl-CoA, which is then dehydrated into trans-enoyl-CoA. The enzymes involved in the two steps are β-ketoacyl-CoA reductase (KCR) and β-hydroxyacyl-CoA dehydratase (HCD), respectively. The last step is the reduction of the enoyl-CoA to generate an acyl-CoA with elongated chain length. This step is catalyzed by enoyl-CoA reductase (ECR).  Figure 1.2 Reactions and enzymes involved in the very-long-chain fatty acid elongation cycle. 10   The substrate specificity of FAE is thought to be determined by the KCS.11 Twenty-one KCS genes have been annotated in the Arabidopsis genome, and they were divided into 4  eight subclasses.11a, 12 Three KCSs from the gene family were found to be involved in cuticle formation: CER6, KCS1 and FDH. In Arabidopsis loss-of-function mutants of CER6 or KCS1, wax compounds with C26 or longer chain lengths were decreased and at the same time an accumulation of C24 wax compounds was observed, suggesting a role of CER6 and KCS1 in the elongation of C24 fatty acyl-CoA. 13 Further evidence came from the heterologous expression experiment of CER6 in yeast, which led to the production of fatty acids up to C28 in chain length. 14 It has been demonstrated recently that the elongation of C28 acyl-CoA to C30 acyl-CoA required the functions of both KCS (CER6) and an additional enzyme: CER2.15 Only when CER2 was co-expressed in yeast with CER6, the C30 fatty acid product could be detected.  Besides KCSs, the KCR, HCD and ECR genes have also been found and characterized in plants. Plant KCRs were first discovered in maize, GL8A and GL8B.16 Their homologous Arabidopsis gene At1g67730 has been found to functionally complement a KCR-deficient yeast mutant.17 CER10 in Arabidopsis has  been identified as the ECR.18 When it was expressed in the yeast mutant lacking ECR activity, the wild-type phenotype of yeast was restored. The discovery of plant HCD was very recent. Two Arabidopsis genes PAS2 and PAS2-1, were found to be homologues of the yeast HCD:PHS1.19 The phenotype of yeast phs1 mutant could be restored by PAS2, and conversely PHS1 could complement the pas2-1 mutant. The enzyme’s substrate 3-hydroxy-acyl-CoA was also detected in the pas2 mutant. Unlike KCSs, KCR, HCD and ECR are thought to have broad compatibilities for substrate chain lengths.20  1.3 The biosynthesis of mono-functionalized wax compounds VLC fatty primary alcohols, alkanes, acids, aldehydes and esters are the major compounds of most plant wax mixtures.21 Primary alcohols, acids and aldehydes are dominated by even-numbered homologues, whereas for alkanes, predominant homologues are of odd-numbered chain lengths. The origin of alkanes has been described as wax 5  compounds that have lost the terminal functional groups. Therefore, even though strictly speaking alkanes do not have functional groups, they can still be regarded as derivatives of mono-functionalized wax compounds.  It is generally believed that after fatty acid elongation wax biosynthesis diverges into the primary alcohol and alkane pathways (Figure 1.3). The first pathway is responsible for converting VLCFAs (CoA thioesters) into primary alcohols and subsequently esters, whereas the second pathway gives rise to aldehydes and alkanes. The two pathways will be discussed separately in the following subsections (1.3.1 and 1.3.2).  Figure 1.3 Biosynthesis of mono-functionalized wax compounds and secondary alcohols and ketones. The VLC acyl-CoAs are the substrates for both the primary alcohol pathway and the alkane pathway. Primary alcohols and esters are produced in the primary alcohol pathway, whereas aldehydes, alkanes, secondary alcohols and ketones are produced in the alkane pathway.  1.3.1 The primary alcohol pathway The first fatty acyl reduction activity was identified from Brassica oleracea and a two-step reduction mechanism was proposed, with aldehyde as an isolable intermediate.22 Then later, a new fatty-acyl reductase (FAR) from jojoba (Simmondsia chinensis) was found able 6  to catalyze the two reductions without releasing the aldehyde intermediate.23 Eight FAR-like genes were identified in Arabidopsis, among which CER4 was characterized to be the FAR involved in the wax biosynthesis.24 The stem wax of cer4 mutants showed a drastic decrease in primary alcohols and esters, while the amounts of alkanes, ketones and other aliphatic compounds were not affected.21, 25 Conversely, when CER4 was expressed in yeast, an accumulation of C24 and C26 alcohols was detected. 24 Subcellular localization studies into the GFP-tagged CER4 protein revealed that it was localized to ER membranes in the epidermal cells of leaves and stems.24 All these evidences were consistent with its role as a FAR in cuticular alcohol biosynthesis.  The resulting primary alcohols are used as substrates to synthesize wax esters. Eleven genes in Arabidopsis were annotated as wax synthase/diacylglycerol acyl transferase (WS/DGAT) bifunctional genes.26 However, only one of them, WSD1 has been cloned and characterized.27 Highest expression of WSD1 was observed in the stem epidermis.28 The stem wax of wsd1 mutants exhibited almost total depletion of alkyl esters, whereas the other compound classes were not affected.27 The wax synthase activity of WSD1 was confirmed by both in vitro and in vivo assays, as opposed to very weak DGAT activity. In the epidermal cells, WSD1 was revealed to localize to the ER, consistent with the assigned function.  1.3.2 Biosynthesis of alkanes Alternative to the alcohol pathway, VLCFAs (or CoAs) can be channeled to the alkane pathway to produce odd-numbered alkanes, and subsequently secondary alcohols and ketones (Figure 1.3). The wax compounds synthesized in the alkane pathway account for 70% to 80% of total wax in Arabidopsis.21, 29 Since alkanes are categorized along with mono-functionalized compounds in this thesis, secondary alcohols and ketones will be regarded as bifunctional wax compounds and their biosynthesis will be discussed in section 1.4.1. 7   Alkanes are widely thought to be biosynthesized via decarbonylation of the aldehyde intermediates. This hypothesis was first proposed based on the findings in pea (Pisum sativum) and green algae (Botryococcus braunii), that C18 acid can be transformed into C17 alkane with the release of CO.30 Indeed, similar two-step pathways have been characterized recently in some cyanobacteria and housefly. In cyanobacteria, the acyl-ACP precursor is first reduced to aldehyde, and then the terminal carbonyl group of C18 aldehyde is cleaved to generate C17 alkane and formate. 31 The decarbonylase belonged to a family of soluble non-heme di-iron enzymes, and it required an external reducing system. In housefly, a P450 oxidative decarbonylase was also found and characterized.32 Similarly, a reducing system (cytochrome P450 reductase + NADPH) was also required by the decarbonylase to produce long-chain alkanes.  In Arabidopsis, two genes, CER1 and CER3, are thought to participate in the biosynthesis of alkanes. First evidence came from analyses of respective mutant waxes. The stem wax of cer3 showed a drastic reduction in all alkane pathway compounds, but an increase in C30 primary alcohol.21 In the cer1 mutant, the stem wax had a dramatic decrease in all alkanes (especially C29) and alkane-derived compounds, but a slightly increased level of C30 aldehyde.21 In addition, a CER1 overexpression line exhibited a notable increase of odd-numbered alkanes in both stem and rosette leaf waxes. These results have led to the proposal that CER3 is the acyl-CoA reductase, which converts acyl-CoAs directly to aldehydes, and CER1 the decarbonylase. This hypothesis was further supported by the subcellular localization of both CER1 and CER3 in the endoplasmic reticulum membranes, where wax biosynthesis takes place.33  Beside phenotype evidence, transmembrane domains have been found in the amino acid sequences of both CER1 and CER3, consistent with their localization to ER.34 In addition, a short sequence containing three histidine-rich motifs HX3H+HX2HH+HX2HH were present in both enzymes.34 The histidine-rich clusters were found to be conserved among a large 8  family of integral membrane enzymes, and they were closely associated with the di-iron catalytic cores.35 The di-iron center in cyanobacterial decarbonylase was critical to the enzyme’s activity, according to the proposed reaction mechanisms.31c, 31e Therefore, it is reasonable to assume that both CER1 and CER3 also participate in the decarbonylation reaction in alkane biosynthesis of Arabidopsis. However, it should be noted that this hypothesis is in contradiction to the earlier model in which the two enzymes catalyzed two different reactions (acyl-CoA reduction and decarbonylation).  Direct evidence for the involvement of CER1 and CER3 in the alkane pathway was provided recently by heterologous expression experiments in yeast.33c In a yeast strain engineered to produce C30 fatty acid substrate, co-expression of CER1 and CER3 resulted in the production of C29 alkane. No alkane was detected if either gene was omitted. The amount of alkane could be greatly increased if CYTB5-B, an electron transferring protein localized in ER, was also expressed in yeast, suggesting a reducing component was required for Arabidopsis alkane biosynthesis. This finding was consistent with the reducing systems found in the characterized decarbonylases. In order to study the functions of histidine-rich clusters in CER1 and CER3, one histidine in each histidine-rich motif was mutated to alanine. The mutated CER1s were not able to reconstitute alkane biosynthesis in yeast, neither could they restore the phenotype of cer1 plants. However, the function of mutated CER3s was the same as the wild-type protein. These results indicated that the histidine motifs were essential for the function of CER1, but not for CER3. Hence CER1 is very likely the decarbonylase in Arabidopsis. More biochemical evidence is still needed to finally identify the substrate and function of both enzymes.  1.4 Biosynthesis of wax compounds with multiple functional groups In addition to wax compounds with functional groups on primary carbons, wax constituents with secondary functionalities have also been discovered from various plant species. In very few cases, these wax compounds are the major or dominant components in cuticular 9  waxes. More commonly, they are the minor or even trace wax constituents. They occur in plant cuticles of various organs and species, and therefore may be very important for the functional diversity of different cuticles.  Secondary functional groups, in the context of wax compounds, are hydroxyl groups and carbonyl groups. Different combinations of secondary/secondary or primary/secondary functional groups can result in diols, ketols, diketones and hydroxyaldehydes (Figure 1.4). The functional groups can scatter along the aliphatic chain, or concentrate in the central or the terminus regions. Hence a large number of positional isomers can be generated.  Figure 1.4 Examples of multifunctional wax compounds  Based on the types and positions of functional groups, the characterized bi- and multifunctional wax constituents can be categorized into four groups: (1) symmetric secondary alcohols, ketones, ketols and diols; (2) β-diketones and hydroxy-β-diketones; (3) primary/secondary bifunctional wax compounds; (4) asymmetric secondary alcohols, ketones and diols. As illustrated by M. Wen in her doctoral thesis, the symmetric compounds are those with functional groups on (or very close to) the central carbons, for example nonacosan-15-ol and 15-hydroxyonacosan-14-one.36 In contrast, the functional groups of asymmetric compounds reside on or towards the terminus of the aliphatic chain, e.g., the secondary hydroxyl group in 1,3-alkanediols.  Except for the symmetric secondary alcohols and ketones, the biosynthesis of secondary 10  functional groups is largely unknown. However, based on available evidence, it has been proposed that the secondary functional groups may be introduced either via hydroxylation (catalyzed by P450 enzymes) or that it may result from the elongation (similar to polyketide biosynthesis). The two pathways have been used to explain the biosynthesis of the bi- and multifunctional wax compounds mentioned above. Each compound group will be discussed separately in the following subsections.  1.4.1 Symmetric secondary alcohols, ketones, ketols and diols Secondary alcohols have been found in waxes of plant species that contain large amounts of alkanes, such as Arabidopsis thaliana, rapeseed and broccoli (Brassica napus and B. oleracea), and pea (Pisum sativum). In the leaf wax of broccoli, nonacosan-15-ol and nonacosan-14-ol have been discovered.37 The stem wax of Arabidopsis contained 30% nonacosan-15-ol and nonacosan-14-ol, along with trace amounts of other isomers. In addition to alcohols, symmetric ketones were also found in the leaf wax of broccoli (Brassica napus) and the stem wax of Arabidopsis.38 In both species, secondary ketones were dominated by nonacosan-15-one.  The fact that symmetric secondary alcohols and ketones were discovered from alkane-rich plant species and their matching chain length profiles had led to the suggestion that both the alcohols and ketones were derived from the corresponding alkane precursors. Direct evidence supporting this hypothesis was provided by early biochemical experiments in which isotope-labeled C29 alkane was transformed into labeled nonacosan-15-ol and nonacosan-15-one by the leaves of broccoli.39 Recently, a reverse genetic approach has led to the isolation of a cytochrome P450-dependent enzyme, mid-chain alkane hydroxylase (MAH1) in Arabidopsis.40 The mah1 mutant lines showed almost complete depletion or greatly reduced levels of secondary alcohols and ketones. Leaves of wild-type Arabidopsis contain only trace amounts of secondary alcohols and ketones. However, they were found in the leaves of MAH1 overexpressor lines. Subcellular localization of MAH1 to 11  the ER membrane was also consistent with its assigned role in wax biosynthesis.40 Therefore, it was concluded that MAH1 was able to synthesize both the symmetric secondary alcohols and ketones from alkanes (Figure 1.3). The hydroxylation reaction is of limited regioselectivity, as it can occur on both C-14 and C-15.  Another group of symmetric wax compounds are the mid-chain diols and ketols. Small amounts of α- and β-ketols were found in the cuticular waxes of broccoli and in the stem wax of Arabidopsis, such as 14-hydroxynonacosan-15-one and 13-hydroxynonacosan-15-one.41  The biosynthetic pathways leading to symmetric ketols and diols are currently unknown. However, it has been proposed that they are synthesized from the symmetric secondary alcohols and ketones (Figure 1.4).41a This is indirectly supported by the fact that the ketols and diols are usually co-occurring with relatively large amounts of symmetric secondary alcohols and ketones, and the positions of functional groups are also very similar between these compound classes. Further evidence came from the stem wax of the mah1 mutant, which was devoid of ketols and diols, together with reduced levels of secondary alcohols and ketones.41b However, in the leaf wax of a MAH1 overexpressor line, where the production of secondary alcohols and ketones was enhanced, no ketols or diols could be detected.41b Therefore, it is likely that a mid-chain-hydroxylase other than MAH1 is responsible for the synthesis of symmetric ketols and diols. Further experiments are required to test this hypothesis. 12   Figure 1.5 The hypothetic pathway leading to symmetric diols and ketols. The symmetric diols and ketols can be biosynthesized from the symmetric secondary alcohols and ketones by a MAH1-like hydroxylase.  1.4.2 β-Diketones and hydroxy-β-diketones Wax β-diketones and hydroxy-β-diketones are characterized by their dominant odd-numbered chain lengths and an in-chain 1,3-diketone motif. The most frequently detected compound in this class is hentriacontane-14,16-dione. It has been found as the only or major diketone compound in barley (Hordeum vulgare)42 and some other species of Gramineae,43 Besides that, other diketone isomers were also found in various plant species, such as nonacosane-8,10-dione44 and hentriacontane-8,10-dione.45 Hydroxy-β-diketones have also been found, usually together with the corresponding diketones, such as 8- and 9-hydroxyhentriacontane-14,16-diones in the leaf wax of wheat,43b suggesting a possible precursor-product relationship.  Based on the positions of functional groups and a direct analogy to polyketide biosynthesis, the introduction of the two carbonyl groups has been proposed to occur during fatty acid elongation (Figure 1.5). By omitting the function of KCR, HCD and ECR in two consecutive elongation cycles, two carbonyl groups can be introduced into the aliphatic chain. This hypothesis can be supported by the results of some biochemical experiments. In barley spikes, inhibitors were found to have different effects on the incorporation of [2-14C] acetate into hydrocarbons and β-diketones.46 Dithiothreitol and mercaptoethanol selectively inhibited the incorporation of labeled acetate into hydrocarbons, whereas arsenite had the 13  opposite effect. Cyanide blocked the synthesis of β-diketones and at the same time increased the production of labeled hydrocarbons. These results suggested the existence of two elongation systems, which indirectly supported the hypothesis that β-diketones resulted from polyketide-like elongation rather than fatty acid elongation. Further evidence was provided by feeding assays with different fatty acids.46 Labeled C16 and shorter chain fatty acids were incorporated into β-diketones, whereas C18 and longer substrates were not accepted. In contrast to that, the chain lengths of fatty acids were found to have no effect on the isotope incorporation into hydrocarbons. This was direct evidence indicating that the biosynthesis of the β-diketone motif starts during the elongation of C16 acyl-CoA to C18 acyl-CoA.   Figure 1.6 The hypothetic pathway leading to hentriacontane-14,16-dione. The β-diketone motif can be introduced by omitting the activities of KCR, HCD and ECR in two consecutive elongation cycles.  1.4.3 Primary/secondary bifunctional wax compounds Unlike the multifunctional wax compounds described above, the primary/secondary bifunctional compounds have even-numbered chain lengths. The secondary functional groups are predominantly found in odd-numbered positions, forming the 1,3-, 1,5-, 1,7-, etc., substitution patterns. Different combinations of the hydroxyl and carbonyl groups on primary and secondary carbons give rise to diols, ketols, hydroxyaldehydes and ketoaldehydes. 14   As illustrated by Wen, These bifunctional wax compounds can be categorized into three groups: with fixed substitution position but varying chain lengths; with fixed chain length and varying positions of secondary functional group; with both varying chain lengths and substitution patterns.36 Examples for each group were found in plants, such as the 1,5-diols (from C28 to C38) and 5-hydroxyaldehydes (from C24 to C36) found in the wax of Taxus baccata needles,47 and the dotriacontane-1,9-diol, -1,11-diol and -1,13-diol from the leaf wax of Myricaria germanica,48  So far, no evidence regarding the biosynthesis of these wax compounds has been reported. However, corresponding biosynthetic pathways can still be hypothesized based on their structures. The strict geometry of secondary functional groups being on the odd-numbered carbons is very similar to that of some polyketide pathways, suggesting the secondary functional groups are likely introduced during polyketide-like acyl elongation.  Taking 1,5-ketol as an example, the biosynthesis of the prim./sec. bifunctional compounds can be illustrated. The carbonyl group may first be introduced by omitting the KCR, HCD and ECR activities in one specific elongation cycle, similar to the biosynthesis of β-diketones. The resulting β-keto-acyl-CoA may undergo one more elongation cycle to afford 5-keto-acyl-CoA. The thioester functionality on the terminal carbon may then be reduced into an alcohol via the primary alcohol pathway, giving the final 1,5-ketol product with even-numbered chain length. The secondary carbonyl group can be further reduced to produce 1,5-diol. Alternatively, the secondary hydroxyl group can also be introduced during elongation by omitting only the HCD and ECR activities. To test these pathway hypotheses, experimental evidence needs to be provided.  1.4.4 Asymmetric secondary alcohols, ketones and secondary/secondary diols Similar to the symmetric secondary alcohols and ketones, also the asymmetric isomers are 15  dominated by odd-numbered chain lengths. However, the functional groups, instead of being located in the center of the aliphatic chain, reside on even-numbered carbons towards the chain terminus, such as nonacosan-10-ol.49 Similar to the primary/secondary bifunctional compounds, the asymmetric secondary alcohols and ketones can be grouped according to chain length profile and substitution positions. Compounds with fixed chain lengths, such as heptacosan-8-ol, -10-ol and -12-ol in the capsule waxes of Papaveraceae,49a as well as those with fixed position of functional group, such as 10-ketones ranging from C27 to C33 in the wax of Osmunda regalis fronds, 50 have all been isolated and characterized. In addition, asymmetric secondary/secondary diols were also detected, usually co-existing with asymmetric secondary alcohols.48-49  The pathways leading to asymmetric secondary alcohols and ketones are poorly understood. However, it is possible that a hydroxylase similar to MAH1 but with different regioselectivity is involved. Primary functional groups such as acid and carbonyl groups may assist regioselectivity. After the secondary functional group is installed, the primary functionality might be cleaved off (via an alkane-like pathway) to afford the odd-numbered aliphatic chain. Another possibility is that the secondary functionality be introduced during elongation, analogous to primary/secondary bifunctional compounds. Both hypotheses are possible at this point, as no evidence is available to rule out any one of them. The biosynthesis of asymmetric diols is possibly derived from the corresponding secondary alcohols, due to their co-occurrence. The second hydroxyl group cannot be introduced by elongation, because each elongation cycle will extend the aliphatic chain by two carbons, therefore the functional group can only be located on alternating carbons. This leaves hydroxylation the only possible choice. Further investigations are required to identify the hydroxylase and to study its regioselectivity.  1.5 Objectives As reviewed in previous sections, wax compounds with primary functional groups have 16  been detected in various plant species, and much progress has been made in understanding their biosynthesis pathways. In contrast to that, relatively little is known about the biosynthesis of the bi- and multifunctional wax compounds, except for the symmetric secondary alcohols and ketones. In order to understand the biosynthesis of these large groups of wax compounds, various combinations of functional groups and chain length ranges need to be first identified from plant waxes. Therefore, the main focus of this study was on the identification of one set of bifunctional compounds. Special focus was on 1,2- and 1,3-bifunctional wax compounds from certain plant species.  The first research objective was to identify the structures of the unknown compounds previously found in Cosmos bipinnatus petal wax (Chapter 3). The unknown compounds had previously been assigned structures with 1,2-bifunctionality and, hence, candidate 1,2-bifunctional standard compounds needed to be synthesized first. Second, two NMR-based approaches were used to complete the structural characterization by studying the stereoconfigurations of the 1,2-alkanediols found in Cosmos bipinnatus petal wax (Chapter 4). The research goal in Chapter 5 was to provide a series of very-long-chain 1,3-bifunctional standard compounds by chemical synthesis, which could be used to search for 1,3-bifunctional wax compounds in plants, such as Aloe arborescens.  Another focus of this study was on the alkane biosynthesis pathway, as no final evidence is available to assign the substrates and functions of the involved enzymes, CER1 and CER3. To accomplish this goal, there were two objectives in this study. The first was to produce isotope-labeled very-long-chain substrates that could be used in future in vivo assays. The second was to synthesize C30 aldehyde as the most likely substrate for CER1, and to test it in assays using transgenic yeast cells.     17  2 Materials and methods 2.1 Plant materials and wax extraction Cosmos bipinnatus was grown under the conditions reported in Christopher Buschhaus’ doctoral thesis51 and Aloe arborescens was grown in a greenhouse under ambient conditions. To extract the total wax from the cosmos petals and aloe leaves, the plant materials were submerged in chloroform for 30 sec. The same extraction process was repeated using fresh chloroform. The organic solutions from the two extractions were combined and the organic solvent was evaporated under a gentle stream of nitrogen gas at 50°C.  2.2 Sample derivatization Proper amounts of wax samples were transferred into sample vials and derivatized with excess bis-N,O-(trimethylsilyl)trifluoroacetamide (BSTFA) and pyridine (1:1, v/v) for 30-40 min at 75°C. Then the excess derivatization agents were evaporated under nitrogen gas flow. Fresh chloroform was added to the vials when the samples were dry. The aloe wax samples were derivatized directly, whereas the cosmos petal wax samples were first separated by TLC (silica gel) using the mobile phase conditions mentioned in Chapter 3 and 4. TLC plates were sprayed with diluted primuline (acetone/H2O) and visualized under 365 nm UV light. Bands of interest were removed from the TLC plates and extracted with fresh chloroform. The extracted solutions were dried under nitrogen gas flow and derivatized as described above.  2.3 GC-MS/FID analysis The prepared wax samples were analyzed by capillary GC (6890N, Agilent, Avondale, PA; column 30 m HP-1, 0.32 mm i.d., df=0.1 μm) using the following temperature program: on-column injection at 50°C, held for 2 min at 50°C, raised by 40°C min-1 to 200°C, held for 2 min at 200°C, raised by 3°C min-1 to 320°C, and held for 30 min at 320°C. For the 18  purpose of elucidating compound structures, the GC was connected to a mass spectrometer (5973N, Agilent, 70 eV electron impact) with a programmed helium carrier gas flow of 1.4 ml min-1. The structures of wax compounds were identified by comparing their mass spectra with those of standard compounds. For quantitative purposes, the GC was connected to a flame ionization detector (FID) with a programmed hydrogen carrier gas flow of 2.0 ml min-1. The relative amounts of wax compounds were calculated based on their peak areas.  2.4 Chemical synthesis All reagents and solvents were commercially purchased and used directly without purification except for THF and CH2Cl2, which were distilled from Na/benzophenone and CaH2, respectively. All reactions were performed in oven-dried flasks without inert atmosphere protection. Analytical TLC was performed on silica gel coated aluminium sheets (layer thickness 0.2 mm). Crude products were purified by preparative TLC plates (silica gel, layer thickness 0.5 - 2 mm, 20*20 cm, with 4 cm of concentration zone) or by flash chromatography (pore size 60 Å, 230−400 mesh). TLC plates were visualized using the same method as described above. The purities of all purified samples were checked by GC-MS to be no less than 90%. 1H-NMR spectra were recorded from CDCl3 solutions at 25°C on Bruker Avance 300 or Bruker Avance 400dir instruments.  The detailed synthetic protocols and characterizations of products of each chapter are included in the appendices: appendix A (Chapter 3), appendix B (Chapter 4), appendix C (Chapter 5) and appendix D (Chapter 6).  2.5 Yeast feeding experiment and lipid extraction 2.5.1 Yeast expression and feeding conditions Sequenced pESC-CER1 plasmid was transferred into wild-type Saccharomyces cerevisiae strain (W3031A) and cultivated on minimal dropout medium lacking tryptophan and 19  containing glucose at 30°C for 3 days. The presence of the plasmid containing the CER1 gene in each yeast colony was verified by PCR. Recombinant yeast cells from the verified colonies were first grown at 30°C overnight in 5 ml inducing minimal medium lacking tryptophan and supplemented with galactose. Then the 5 ml culture was transferred to 25 ml inducing medium and grown for another 16 hours. Substrate (C30 aldehyde or alcohol, 0.5 mg) in hot ethanol was added to the 25 ml culture, and it was grown for another 24 h before lipid extraction.  2.5.2 Yeast total lipid extraction Yeast cells were separated from culture medium by centrifugation and dried under nitrogen gas flow. Dried yeast cells were then subject to transmethylation with 2 ml methanol and 400 μl at 90°C for 1.5 h. After cooling, 1 ml NaCl 2.5% (w/v) was added and the total lipids were extracted with 2*2 ml of hexane. Extracts were washed with 2 ml NaCl 2.5% (w/v) and then dried under a gentle stream of nitrogen. The sample was derivatized and analyzed by GC-FID/MS as described above. The culture medium was also extracted with 2*10 ml of hexane and analyzed the same way.         20  3 Identification and quantification of 1,2-diol monoacetates in Cosmos bipinnatus petal wax 3.1 Introduction In the on-going project to elucidate the relationship between wax composition and function, previous Ph.D. student Christopher Buschhaus started to investigate the compositions of petal waxes from Cosmos bipinnatus. While major functions of the petal cuticles are to prevent un-controlled water loss and to protect from UV light and other stresses, some additional functions such as scent emission and physical contacts with pollinators may turn out to be equally important. Thus it is very likely that the wax constituents on plant petals may differ from those found on other plant organs.  The ability of C. bipinnatus petal wax to block water was found to be less than half that of leaves.51 The reduced water permeability was likely the result of different chemical composition in the petal waxes. Therefore, a thorough analysis of the wax composition was necessary to establish the function-composition relationships for the petal waxes. Analyses of C. bipinnatus petal waxes revealed some unusual features,51 including (1) a total wax load of only 25%–33% of that on leaves or stems; (2) a primary alcohol concentration of more than 50% in the total wax mixture; (3) C22 and C24 as the predominant chain lengths, as opposed to C30 compounds in stems and leaves. In addition, unknown compounds were discovered in two TLC fractions during the analysis of wax samples extracted from C. bipinnatus petals. Since those unknown compounds may be a crucial clue for understanding the special functions of petal cuticles, further endeavors were carried out to identify these compounds and quantify their amounts in the petal waxes.  One of the unknown fractions migrated between the origin and the VLC fatty acid band on TLC.51 Two homologous series A1 and A2 were detected in this fraction by GC-MS, with seven compounds in each series. By comparing the GC chromatograms and mass spectra of TMSi-derivatized unknown compounds and  synthetic standards, the compounds in 21  series A1 and A2 were identified as alkane 1,2-diols and 1,3-diols, respectively (Figure 3.1).51-52 The novel 1,2-substitution pattern did not fall into any of the compound groups described previously (see Section 1.4).   Figure 3.1 Mass spectra of diols from C. bipinnatus petal waxes. 51  (A) Bis-TMSi ether of 1,2-docosanediol from series A1; (B) Bis-TMSi ether of 1,3-docosanediol from series A2.  Based on molecular ions and α-fragment patterns, the chain lengths of compounds within both series A1 and A2 were found to range from C20 to C26. Compounds with even carbon number had much higher abundance than odd-chain compounds. The most abundant chain length was C22 for both series, constituting roughly 60% of the 1,2-diols and 72% of the 1,3-diols. 1,2-diols were found to make up 3-5% of the total wax load, and the amount of 1,3-diols was approximately one order of maganitude less.  The other cosmos petal wax fraction containing unknown compounds migrated between free acids and primary alcohols on TLC.51 GC-MS analysis revealed two series of compounds in this fraction: B1 and B2. Series B1 contained seven homologous compounds, while B2 consisted of five homologues. Reaction of the fraction with lithium A B 22  aluminum hydride yielded a series of 1,2-diol homologues, showing that both B1 and B2 were composed of novel bifunctional compounds with the functional groups on primary and adjacent secondary carbons. Because, based on their TLC behavior, the unknowns were less polar than fatty acids and more polar than alcohols, there could only be one hydroxyl group in the B1 and B2 compounds. This conclusion was further supported by the m/z 73 ion [OTMS]+ in the mass spectra of the TMSi-derivatized unknowns (Figure 3.2). It was also concluded, based on the abundance of ion m/z 103 [CH2OTMSi] + in the mass spectra, that the hydroxyl groups were at the primary positions of the compounds in series B1, and in secondary positions in B2. Based on polarities, the other functional group could be a carbonyl or a protected alcohol (such as an ester).  Figure 3.2 Mass spectra of unknowns in series B1 and B2. 51  (A) TMSi ether of the most abundant compound in series B1; (B) TMSi ether of the most abundant compound in series B2. The total amount of unknown compounds was found to be 0.04 ± 0.01 μg/cm2 on the adaxial (upper) side and 0.09 ± 0.01 μg/cm2 on the abaxial side (lower surfaces) of the petal, respectively. The compounds thus constituted 1.8 ± 0.4% and 3.3 ± 0.2% of the adaxial and abaxial wax mixtures, respectively.  All the previous evidence taken together, the most likely structures for the unknowns in B1 B A 23  and B2 were either 1-hydroxy-2-ketone and 2-hydroxy aldehyde, or else esterified 1,2-diols. The goal of this project was to structurally identify and quantify the unknowns in B1 and B2.  3.2 Results To identify the unknown compounds, standards were synthesized for proposed candidate structures. They were characterized by GC-MS and 1H-NMR and compared to the spectral and chromatographic characteristics of the unknowns (section 3.2.1 and 3.2.2). To further distinguish isomeric structures and test for interconversion between them, possible acyl-migration reactions were further studied (section 3.2.3). Finally the relative amounts of the two unknown series in the petal waxes could be quantified (section 3.2.4).  3.2.1 Synthesis of alkane 1-hydroxy-alkane-2-one standards The first goal was to synthesize an α-hydroxy ketone standard, as a candidate structure for the unknowns in B1. α-Hydroxy ketones are important intermediates in organic synthesis. Common strategies to synthesize α-hydroxy ketones include α-hydroxylation of carbonyl compounds,53 selective oxidation of vicinal diols54, or selective oxidation of alkenes.55 Various oxidation reagents and methods have been developed to directly oxidize vicinal diols into hydroxyketones, such as dimethyldioxirane or its trifluoromethyl analogue,56 the H2O2/TS-1 system, 57 NaBrO3/NaHSO3 58 and electro-chemical methods.59 Another approach is to convert the diols into their cyclic stannylene derivatives. The O-Sn bonds formed have distinct stabilities and turn into active secondary rather than primary O-Sn bonds.60 Examples have been reported using the stannylene derivatives of diols to selectively oxidize secondary alcohols in carbohydrates,61 cyclic62 and aliphatic diols63 with satisfactory yields. Here the stannylene derivative approach was chosen for the synthesis of the 1-hydroxy-2-ketone standards due to its broad versatility with diol substrates and reliable conversion yields.  First, reaction conditions were tested on a short-chain vicinal diol. 1,2-Octanediol was 24  converted into its cyclic dibutylstannylene derivative 3-1 by refluxing with dibutyltin oxide in methanol. The conversion was almost quantitative. Brominolysis of the stannylene ether selectively afforded product 3-3 with oxidized secondary hydroxyl, with a two-step yield of 74%. The homologous 1-hydroxy ketone 3-4 was synthesized from 1,2-tetradecanediol using the same reaction conditions with an overall yield of 78%. Both compounds were analyzed by 1H-NMR, and the signals matched those of similar structures reported in the literature.55, 63  Scheme 3.1 Synthesis of 1-hydroxy-2-ketone standards.  The standards were TMSi-derivatized and then subjected to GC-MS analysis (Figure 3.3). The resulting mass spectra of both compounds showed ions at m/z 73 and 103 as signature for primary alcohols and also ions at m/z 117, 129 and 143 that cannot be assigned directly. The base peaks were due to loss of methyl groups [M-15] +, and α-fragments were also detected at m/z 113 [C6H13CO] + and m/z 197 [C12H25CO] + for the C8 and the C14 products, respectively.  25   Figure 3.3 Mass spectra of 1-hydroxy-alkan-2-one standards. (A) TMSi ether of 1-hydroxyoctan-2-one; (B) TMSi ether of 1-hydroxytetradecan-2-one. Comparing the mass spectra of the 1-hydroxy-alkan-2-one standards to those of the unknowns in series B1 and B2 (Figure 3.2, A), obvious differences were observed. While all unknowns and standard spectra had ions with m/z 73, 103, 117 and 129, their relative abundances were quite different. In the mass spectra of B1, fragment m/z 117 was the base peak. In the mass spectra of B2, fragment m/z 73 had much lower intensity and fragment m/z 103 was barely detectable. In addition, a pair of peaks with similar intensities and a difference of 15 mass units were signature to the unknowns in series B1. However, this fragmentation pattern was missing in the mass spectra of 1-hydroxy-alkan-2-one standards. Hence, it was concluded that the unknowns in series B1 and B2 were not 1-hydroxy-2-ketones. The mass spectra of B1 and B2 had implied that the structures of both unknown series might be very similar. Therefore, after 1-hydroxyketone structures had been ruled out, it also appeared unlikely that one of the series could comprise 2-hydroxyaldehydes.  3.2.2 Synthesis of 1,2-diol monoacetates standards The next goal was to synthesize the other possible structures for the unknown compounds: esterified 1,2-diols, as described above. The standards were first synthesized using the A B 26  commercially available 1,2-tetradecanediol as starting material. Reacting this compound with an equivalent amount of acetic anhydride in the presence of pyridine was expected to produce a mixture of 1,2-diol acetates (Scheme 3.2). The resulting mixture was derivatized with BSTFA and then analyzed by GC-MS. Four compounds were detected (Figure 3.4) and the peak with shortest retention time (13.01 min) was identified as 1,2-tetradecanediol bis-TMSi ether. To identify the others, 1,2-tetradecanediol bis-acetate was synthesized using the same protocol but with large excess of acetic anhydride and subjected to analysis on GC-MS. The peak with longest retention time (13.28 min) was thus identified as 1,2-tetradecanediol bis-acetate. This left the two peaks in the middle for 2-hydroxytetradecyl acetate (3-5) and 1-hydroxytetradec-2-yl acetate (3-6). Scheme 3.2 Synthesis of 1, 2-diol-monoacetate standards.   Figure 3.4 GC-MS chromatogram of the product mixture from the acetylation of tetradecane-1,2-diol.  In order to compare the acetates with the unknowns, the mass spectra of the two synthetic monoacetates were carefully examined (Figure 3.5). The peak at 13.15 min exhibited a MS fragmentation pattern of m/z 117 as base peak, twin peaks at m/z [M-60]+ (284) and [M-75]+ 27  (269) and the fragments m/z 73, 103, 129, all very similar to the fragmentation pattern of the unknowns in B1 (see Figure 3.2). Because the fragment m/z 103 was indicative for a primary alcohol, this peak was tentatively assigned as 1-hydroxytetradec-2-yl acetate 3-6. Accordingly, the only peak left (13.22 min) was assigned as 2-hydroxytetradecyl acetate 3-5. Its base peak was at m/z [M-73]+ (271), accompanied by ions m/z 73, 117, and 175. All fragments together suggested structural similarities to the unknowns in B2.  Figure 3.5 Mass spectra of acetates synthesized from 1,2-tetradecanediol.  (A) TMSi ether of 1-hydroxytetradec-2-yl acetate; (B) TMSi ether of 2-hydroxytetradecyl acetate.  To further confirm the positions of acetyl groups in each standard, efforts were made to synthesize the 2-hydroxytetradecyl acetate 3-5 from the previously obtained α-hydroxy ketone 3-4 (Scheme 3.3). The free alcohol group in 3-4 was derivatized with acetic anhydride and pyridine to afford 2-oxotetradecyl acetate 3-9, the structure of which was confirmed by 1H-NMR. Further reaction of 3-9 with NaBH4 was expected to result in the reduction of the ketone 3-5. Quite unexpectedly, when the isolated product was analyzed by GC-MS and 1H-NMR, it turned out to be a mixture of monoacetates, with relative amounts of 3-5 and 3-6 of roughly 2:1. The formation of 3-6 suggested that the acetyl group migrated to the secondary hydroxyl group after the reduction had been completed. B A 28  This phenomenon will be discussed in more detail in section 3.2.3. Scheme 3.3 Synthesis of 2-hydroxytetradecyl acetate from 3-4. The next step was to separate the two monoacetates and analyze them individually on NMR. Various TLC mobile phase systems were tested for their abilities to separate the two isomers. In pure chloroform, dichloromethane and acetonitrile, the two isomers co-migrated. Separation was achieved in a mixture of acetone and hexane (2:5), with the Rf values of 0.53 and 0.48 for the two isomers 3-5 and 3-6, respectively. Similarly, in ethyl acetate and hexane (1:3), Rf values were 0.59 for 3-5 and 0.49 for 3-6. The best separation was achieved using pure 1,1,1-trichrloroethane, where 3-5 migrated to 0.48 and 3-6 to 0.35, with a ΔRf value of 0.13.  The isolated 1-hydroxytetradec-2-yl acetate and 2-hydroxytetradecyl acetate were subjected to 1H-NMR analysis (Figure 3.6). In general, the proton adjacent to a hydroxyl (-OH) group resides in higher field in NMR than the α-proton of an acetyl group (-OCOCH3). This pattern was indeed observed in the NMR spectra of the two isomers. The protons on the secondary carbon of the tentatively assigned 2-hydroxytetradecyl acetate 3-5 had much smaller chemical shift (3.84 ppm) than that of 1-hydroxytetradec-2-yl acetate 3-6 (4.90 ppm). On the other hand, the primary protons from 3-5, being next to an acetyl group, were in lower field (4.15 ppm, 3.95 ppm) when compared to the primary protons from 3-6 (3.72 ppm. 3.63 ppm). It was also observed that the methyl protons (-OCOCH3) from the primary acetyl group were in slightly lower field (2.10 ppm) than those in the secondary acetyl group (2.09 ppm). The chemical shifts of all signature peaks mentioned above 29  matched corresponding signals of comparable compounds reported in the literature.64 Thus, the previously assigned structures of 1,2-diol 1- and 2-acetates were confirmed. Mono- Acetates -CH(OR1)CH2OR 2 5.25 – 3.25 ppm -OCOCH3 2.20 – 2.00 ppm    3-5 R 1  = H R 2  = Ac  3-6 R 1  = Ac R 2  = H    Figure 3.6 Characteristic 1 H-NMR signals of isolated 2-hydroxydocosyl acetate and 1-hydroxydocosan-2-yl acetate. To unambiguously identify the structures of the unknowns, standards with similar carbon chain length (C20 to C26) were required. For that purpose 1,2-docosanediol was synthesized from 2-hydroxydocosanoic acid by refluxing with lithium aluminum hydride, and subsequently converted into its monoacetates using similar esterification methods as mentioned above. The resulting mixture was analyzed in the same manner as before. The mass spectra of the 1-hydroxydocosyl acetate 3-8 and 2-hydroxydocosyl acetate 3-7 standards were identical to the mass spectra of the most abundant peaks in B1 and B2, respectively (Figure 3.2).  The two isomeric monoacetates 3-7 and 3-8 both had fragments m/z 441 [M-15]+ (compare Figure 3.2). In the mass spectrum of 3-8, the presence of abundant fragments m/z 73 and 103 indicated a primary hydroxyl function (Scheme 3.4). The signature twin fragment peaks m/z 396 [M-60]+ and m/z 381 [M-60-15]+ can be attributed to loss of the acetyl group and an additional methyl group, respectively. A very small α-fragment was observed at m/z 30  353 [C20H41COAc] +, suggesting the position of the acetyl group. However, the structures of ions at m/z 75, 117, 129, 135 and 306 remain unclear. One possible explanation for the abundant fragment m/z 117 was [CHOCH2OSi(CH3)2] +, but further investigation is required to verify this. In the mass spectrum of 3-7, the most abundant ion at m/z 383 [C21H42OTMSi] + can be explained as the α-fragment due to the cleavage of the bond between C-1 and C-2. The α-fragmentation between C-2 and C-3 would give rise to the ion m/z 175 [CHOTMSiCH2OAc] +. All fragmentation information thus indicated that the position of acetylation in 3-7 was on the primary alcohol, rather than secondary.   Scheme 3.4 Fragmentation patterns of the TMSi ethers of 1,2-docosanediol monoacetates. Further tests showed that the two pairs of compounds had the same retention times during GC separation (Figure 3.7). Thus, the unknown series B1 from cosmos petal wax could be assigned as 1,2-diol 2-acetates, and series B2 assigned as 1,2-diol 1-acetates. 31   Figure 3.7 Co-elution experiment of the TLC fraction that contains 1,2-diol monoacetates and 1,2-docosanediol monoacetate standards. The most abundant peak in series B1 had approximately the same retention time as 3-6, and similarly the most abundant peak in series B2 had almost the same retention time as 3-5.  By examining the chain-length specific fragments of all compounds in the B1 and B2 series, the structure of other homologues could be identified (Figure 3.8). In B1, chain lengths ranged from C20 to C26, while the chain lengths in B2 were from C20 to C24. The even-numbered chain lengths were found to be predominant in both series.        32    Figure 3.8 Mass spectra of two representatives each from series B1 and B2.  (A) Identified as TMSi ether of 1-hydroxyeicosan-2-yl acetate; (B) identified as TMSi ether of 2-hydroxyeicosyl acetate; (C) identified as TMSi ether of 1-hydroxytetracosan-2-yl acetate; (D) identified as TMSi ether of 2-hydroxytetracosyl acetate. For the mass spectra of TMSi derivatized 1-hydroxydocosan-2-yl acetate and 2-hydroxydocosyl acetate, refer to Figure 3.2.  A B D C 33  3.2.3 Study of the acyl-migration behavior of the Cosmos bipinnatus 1,2-diol monoacetates As mentioned above, 3-6 had formed during the synthesis of 2-hydroxytetradecyl acetate 3-5, due to migration of the acyl group from the primary to the secondary alcohol function. This finding raised the question whether both monoacetates were bio-synthesized, or whether only one was produced by plants and the other one formed as an artifact during sample preparation or analysis. To answer this question, the behavior of acetyl groups on the two hydroxyl groups needed to be further investigated.  The behavior of 3-5 and 3-6 was studied with NMR to avoid acyl-migration possibly occurring due to elevated temperatures during derivatization and analysis by GC. In solution (CDCl3) at ambient temperature both compounds were slowly inter-converting (Figure 3.9). After two days in solution, the acyl-migration products had become obvious in both samples. In sample 3-6, approximately at day 5, the amount of acyl-migrated product 3-5 started to surpass that of the starting material of secondary monoacetate. After one week (day 9), the ratios of the two isomers in both samples were roughly the same based on NMR peak integrations, suggesting that the equilibrium had established from both directions. 3-5 was clearly the predominant isomer and thus thermodynamically more stable. The equilibrium ratio of 7:3 between 3-5 and 3-6, respectively, was very close to the ratio observed by GC after reacting the diol with Ac2O and pyridine (see Figure 3.4).         34   –CH(OR 1)CH2OR 2 5.25 – 3.25 ppm –OCOCH3  2.15-2.05 ppm 3-5 R 1  = H R 2  =Ac  3-6 R 1  = Ac R 2  =H  Figure 3.9 Study of acyl migration using 1 H-NMR. Acyl migration was observed from day 1 (D1) to day 9 (D9) for both primary monoacetate (3-5) and secondary monoacetate (3-6) in CDCl3 solution.  Acyl migration due to other factors involved in sample preparation and analysis steps was also studied. In general, the primary monoacetate 3-5 was fairly stable under storage conditions (solid state, -20°C) and on silica gel while purifying by TLC, while slow inter-conversion was detected by NMR for 3-6 under the same conditions. Reaction with only BSTFA caused minor or no acyl migrations for both isomers. However, the addition of pyridine caused substantial inter-conversion of isomers.  3.2.4 Analysis and quantification of 1,2-diol monoacetates in C. bipinnatus petal wax The total amounts of 1,2-diol monoacetates had already been quantified, as mentioned before. However, the ratios between the monoacetate isomers had not been determined.  ppm (t1) 3.504.004.505.00 0 50 100 150 ppm (t1) 2.0502.1002.150 0 500 1000 ppm (t1) 3.504.004.505.00 0 10 20 30 ppm (t1) 2.0502.1002.150 50 100 150 200 250 D2 D5 D9 D1 D2 D5 D9 D1 D1 D2 D5 D9 D1 D2 D5 D9 35  A C. bipinnatus petal wax sample was fractionated by TLC using a mixture of chloroform and ethanol (99:1), and the fraction containing 1,2-diol monoacetates was extracted. After derivatization (with BSTFA alone), the mixture was analyzed with both GC-MS (Figure 3.10) and GC-FID using the same protocols as for previous wax analysis. The structures of all monoacetates were identified by MS, and their amounts were quantified relative to the most abundant compound 1-hydroxydocosan-2-yl acetate (set as 100) by their peak areas on GC-FID. Two independent wax samples were analyzed and results averaged to account for biological variations.   Figure 3.10 GC-MS chromatogram of the TLC fraction containing 1,2-diol-monoacetates. Abbreviations: ac-acid, ol-alcohol, sm-secondary monoacetate, pm-primary monoacetate.  C22 diol monoacetates dominated the C. bipinnatus petal wax fraction, corresponding to 46% of total monoacetates (Figure 3.11). The C24 homologues constituted roughly 1/3 of the total monoacetates. The remaining 25% was C20 diol monoacetates.  36   Figure 3.11 Relative amounts of 1,2-diol monoacetates (% of C22 diol 2-acetate) extracted from C. bipinnatus petal. Results were averaged between two independent samples.  For all chain lengths, the amounts of secondary monoacetates surpassed those of the corresponding primary monoacetates (Table 3.1). The ratios of secondary and primary monoacetates ranged from 4.6: 3 to 5.4: 3, with an overall average of approximately 5:3, opposite to the equilibrium ratio of 3: 7.  Table 3.1 Chain-length specific ratios of secondary monoacetates and primary monoacetates. Results were averaged between two independent samples. C20 diol 2-acetate : C20 diol 1-acetate 5.0 : 3.0 C22 diol 2-acetate : C22 diol 1-acetate 4.6 : 3.0 C24 diol 2-acetate : C24 diol 1-acetate 5.4 : 3.0 Average 5.0 : 3.0  3.3 Discussion The compounds in the two unknown series B1 and B2 in Cosmos bipinnatus petal wax were identified as homologues of 1,2-diol 2-acetates and 1-acetates by comparing the mass spectra and GC retention times with those of synthetic standards. They are novel structures for plants. However in the preorbital secretion of Raphicerus campestris (steenbok), C20 and C22 diol monoacetates had been identified previously. 65 Other ester 37  derivatives of long-chain and very-long-chain aliphatic 1,2-diols had also been reported for waxes of bacteria,66 insects,67 birds68 and mammals.69  Migration of the acetyl group was observed during the synthesis of 3-6. Similar acyl migrations had been reported for other vicinal diols70 and poly-hydroxy natural products, such as glycerols,71 sugars72 and nucleosides.73 An equilibrium ratio of 7:3 between primary and secondary monoacetates was reached from both directions in the present time-course NMR study. Ratios of 6.6:3 and 2:1 were also obtained between primary and secondary monoacetates during the reduction of 3-9 and acetylation of diols, respectively. The approximate ratio of 7:3 matched those found in other 1, 2-diol mono-acyl esters,70a-c, 70f and can therefore be used to describe the thermodynamic equilibrium of the two isomers.  On the plant petals, a ratio of 3:5 of primary to secondary monoacetates was measured. The fact that thermodynamically less favored 1,2-diol 2-acetates were found to be more abundant in the plant wax might suggest that they were preferentially or even exclusively biosynthesized in plant. Conversely, the ratio could also be explained by preferential hydrolysis of the 1-acetate isomer by some esterase. Possible biosynthetic pathways of 1,2-diol monoacetates will be discussed in Chapter 4.  It has long been proposed that the functions of cuticle wax are closely associated to the size, shape and alignment of crystalline domains in the wax coating the plant surface.74 The secondary functional groups on the aliphatic chains may affect the formation of such crystals and therefore increase water permeability. Recent studies have discovered the presence of polar domains in cuticular wax that facilitate the penetration of polar compounds.75 The newly identified 1,2-diols, 1,3-diols and 1,2-diol-monoacetates fall into the category of bifunctional wax compounds. Their special structural characteristics may have a unique effect on the microstructure of wax crystals and thus the function of petal cuticles. For example, the bifunctionality may have a special role in the formation of certain 38  polar domains that facilitate emission of molecules to attract pollinators. Further investigations are required to fully understand the contribution of those bifunctional compounds to wax function.                        39  4 NMR-based determination of the stereoconfigurations of cosmos 1,2-diols 4.1 Introduction Novel compounds with 1,2- or 1,3-bifunctional structures had been identified in Cosmos bipinnatus petal wax based on their polarities, NMR behaviors as well as MS fragmentation patterns (see Chapter 3).52 However, one last piece of information on their exact structure was still missing: the stereoconfiguration of the carbons bearing the secondary hydroxyl function. The stereochemistry may provide insights into the biosynthetic origin of the compounds, as well as their potential functions in petal waxes.  The most commonly used NMR-based method to determine absolute configurations is known as Mosher’s method.76 In 1969, Mosher and his colleagues first used α-methoxy-α-trifluoromethylphenylacetic acid (MTPA) and α-methoxy-α-trifluoromethylphenylacetic acid chloride (MTPA-Cl) to esterify secondary alcohols and then elucidate their stereoconfigurations by NMR (Scheme 4.1).77 Later, introduction of higher-field superconducting magnetic resonance spectrometers further improved this method.78 A model has been used successfully to interpret the experimental results for mono-functional compounds. According to this model, molecules adopt conformations that allow the ester bond, the trifluoromethyl group of MTPA and the methine proton to be co-planar (Scheme 4.1). Due to the anisotropic nature of the aromatic ring, the group above the phenyl group of MTPA will be magnetically shielded, therefore giving NMR signalsshifted toward higher field. By calculating the chemical shift difference of the substituent between the S-MTPA and R-MTPA ester derivatives of the alcohol (ΔδSR=δS-δR), the stereoconfiguration can be determined. If the sign for ΔδSR is positive, then the substituent is in the position of R2, if it is negative then the substituent resides in R1. It should be noted that the acid chloride form of MTPA has the opposite S/R configuration from the parent acid, according to the priority rules of substituents.  40   Scheme 4.1 Chiral derivatization agents and reactions for Mosher’s method.  After Mosher’s ground-breaking discoveries, various other chiral derivatization agents (CDAs) have been found. Some CDAs possess structures similar to Mosher’s reagent, like methoxyphenyl acetic acid (MTA)79 and 2-(anthracen-9-yl)-2-methoxyacetic acid (AMA),80 and other CDAs are designed based on totally different chemical reactions involving boronic acid,81 aldehyde82 and dichlorophosphine.83  In 2006, Bull, James and co-workers developed a three-reagent system to determine the stereoconfigurations of diols and primary amines based on their reaction with 2-formylphenylboronic acid (Scheme 4.2).84 One advantage of this system lies in its broad versatility. With a chiral primary amine reagent, this method can be used to determine the stereochemistry of diols,85 and conversely with a chiral diol reagent the same method can applied to amines.86 Another merit of this method is that in most cases the chemical shifts of protons on R3, R3’, R4, R4’and R5 as well as the one on the N=C bond can all respond to the changes of the chemical and magnetic environment, providing multiple sets of signals for analysis. The stereoconfigurations of several simple and complex diols87 as well as poly-hydroxy systems88 have been analyzed using this method since its publication.  41   Scheme 4.2 Boronic acid-based three-reagent derivatization system.  The goal of this research project was to determine the stereoconfigurations of 1,2-diols found in C. bipinnatus petal wax using NMR-based methods, such as Mosher’s method and the recently developed three-reagent system. 1,3-bifunctional compounds were not included because their amounts had been found to be one order of magnitude lower than those of 1,2-bifunctional compounds in the wax, rendering purification and NMR analysis extremely difficult.  4.2 Results The synthesis of enantiomerically enriched standards and isolation of pure 1,2-diols are shown in section 4.2.1, respectively. The NMR-based methods were first established on standards, and then employed on wax compounds to assign their stereoconfigurations (sections 4.2.2 and 4.2.3).  4.2.1 Synthesis of S- and R-1,2-docosanediol standards and isolation of 1,2-diols from C. bipinnatus petal wax Standards are not necessary for the Mosher’s method, if a model can be used to predict the stereoconfigurations. However, for diols it is impossible to predict their stereoconfigurations based on current models. In addition, there is no available model for the boronic acid-based method. Therefore efforts were made to synthesize the two enantiomerically enriched standards.  To synthesize enantiomerically pure 1,2-diols, two methods are commonly used: Noyori asymmetric hydrogenation89 and Sharpless asymmetric dihydroxylation.90 The second 42  method was chosen here mainly due to its easily-handled reaction conditions, even though the β-keto-ester substrates for Noyori asymmetric hydrogenation would have been available from another project (see Chapter 5).  S- and R-1,2-docosanediol were synthesized from commercially available 1-docosene, following a procedure published for a similar product.91 The yields obtained for both products were low, with 12% for 4-1 and 40% for 4-2. Considerable amounts of starting materials were found unreacted and were recycled. However, further optimization was not necessary for the purpose of this project. All synthesized standards were characterized by 1H-NMR and GC-MS.   Scheme 4.3 Synthesis of S- and R-1,2-docosanediol standards.  In order to use the NMR-based approaches, 1,2-diols with high purity needed to be provided. The Cosmos petal wax was extracted and fractioned by TLC as described before (see 3.2.4). The fraction containing 1,2-diols was further purified by TLC using a mixture of chloroform and ethyl acetate (7:1). NMR analysis of the purified 1,2-diols showed chemical shifts matching those of 1,2-tetradecanediol standard. However, the splitting patterns of the protons adjacent to the primary hydroxyl group were poorly resolved, possibly due to the fact that the sample contained a mixture of diol homologues. Mass spectra of the sample further confirmed their diol structures. The diols were quantified using GC-FID with reference to tetracosane internal standard (Figure 4.1). 3.8 mg of 1,2-diols was obtained from the petal wax with a purity of 87%. 43   Figure 4.1 GC-FID chromatogram of purified 1,2-diols from C. bipinnatus petal wax. Peak abundances were normalized according to C24 alkane internal standard (set as 100).  4.2.2 Stereochemical investigation of 1,2-bifunctional wax compounds using Mosher’s Method Two sets of experiments were done. First the method was established with racemic diol standard. Then it was applied to the enantiomerically enriched standards in comparison with wax compounds (Scheme 4.4).   Scheme 4.4 Synthesis of mono-MTPA-derivatized 1,2-diols.  Racemic 1,2-tetradecanediol was reacted with R-MTPA-Cl and pyridine in dry chloroform overnight (Scheme 4.4, A). The mixture was then separated by TLC, and the major product 44  was analyzed by NMR and GC-MS.  The NMR spectrum of 4-3 confirmed the presence of the MTPA group (Figure 4.2, A). Around 4.3 ppm, two sets of peak doublet-doublets were observed, indicating that the two isomers could be differentiated by this method. However, peak integrations suggested the presence of only one MTPA group per diol functionality. Accordingly, the MS ion m/z 503 [M-15]+ corresponded to 1,2-diol mono-MTPA ester TMSi ether, and the ion at m/z 349 matched the mass of α-fragment [CH(OTMSi)CH2O-MTPA] + (Figure 4.2, B and C). The position of the MTPA group could be deduced from the most abundant fragment m/z 271 [C12H25CHOTMSi] +, which was due to α-cleavage between C-1 and C-2. Hence, the product structure was corroborated as mono-MTPA ester of the primary hydroxyl group. Very likely, the isomeric secondary mono-MTPA ester was also formed during the reaction as the minor product. However, only the major product was isolated from the TLC plate and subjected to NMR analysis.            45      Figure 4.2 Characterization of 1,2-tetradecanediol 1-S-MTPA-ester 4-3. (A) 1 H-NMR spectrum (400 MHz) of 4-3; (B) Mass spectrum and (C) major fragmentations of the TMSi ether of 4-3.  In the NMR spectrum of the racemic sample, four peaks with equal abundance were observed in a doublet of doublets (Figure 4.3). They were assigned to the two methylene protons of the two diastereomers.    A B C 46  Structures -CH(OH)CHHCH-OMTPA 4.0 – 4.5 ppm 4-3 4-4 4-5 Mono-(S)-MTPA ester of 1,2-diols from cosmos petal wax Figure 4.3 Signature 1 H-NMR (400 MHz) signals of the 1,2-diol 1-S-MTPA esters.  The enantiomerically enriched standards 4-1, 4-2 (Scheme 4.4, B and C) were transformed into MTPA derivatives and characterized using the same protocol as described above. GC-MS confirmed that the MTPA was attached to the primary hydroxyl in all samples. In the NMR spectra of 4-4 and 4-5, the methoxy protons and the protons on the chiral carbon showed no differences. The only difference observed was for the methylene protons adjacent to the MTPA ester group (Figure 4.3).  Based on the two enantiomerically enriched samples, the two sets of peaks can be assigned. The most up-field and most down-field dd peaks (H1 = 4.36 ppm, J = 11.4, 3.0 Hz; H2 = 4.19 ppm, J = 11.4, 7.6 Hz) arose from 4-4, and the ones in the middle (H2’ = 4.34 ppm, J = 11.4, 3.2 Hz; H1’ = 4.25 ppm, J = 11.4, 7.0 Hz) can be attributed to its diastereomer 4-5. The enantiomeric purities for both standards could not be accurately calculated due to peak overlap. Further discrimination of the proton sets proved to be H1 H2 H2’ H1’ 47  difficult due to lack of a configurational model for the 1,2-diol mono-MTPA ester needed to explain the NMR results. However, since protons H1 and H2’ had smaller vicinal coupling constants, it could be concluded that they were the protons further away from the β-proton than H2 and H1’. Conversely, the latter two were positioned closer to the β-proton due to larger vicinal coupling constants. H2 and H1’ were both shifted up-field, suggesting that they resided in the shielding zone of the phenyl group.  The same condition was now used to derivatize the diol mixture from cosmos wax. When comparing the resulting NMR spectrum of the 1,2-diol mono-MTPA ester mixture to those of standards, the majority of 1,2-diols was found to possess the same configuration as 4-5, and could therefore be assigned as R-1,2-diols. Small amounts of S-1,2-diols were also found to be present in the mixture. Even though overlapping of the signature peaks made it difficult to unambiguously interpret the peak ratio, attempt was still made to roughly estimate the enantiomeric purity of the wax diols, by calculating the ratios of the integrations of the left doublet peak of H1 and the right doublet peak of H2’ (see Figure 4.3). The peak integration ratio for the racemic sample 4-3 was 1.6:1, and that for the wax diol sample was 3.8:1. Since the racemic sample 4-3 corresponded to a 1:1 ratio of the two diastereomers, the percentage of R-diols in the wax mixture could then be calculated accordingly as 70%.  4.2.3 Study of the stereochemistry of 1,2-bifunctional wax compounds using boronic acid-based method. To test the stereoconfigurations of wax alkanediols with a second independent method, a boronic acid-based approach was used. As above, the feasibility of this method was first tested using racemic 1,2-tetradecanediol (Scheme 4.5). According to previous publications,84b, 85 the three components were expected to afford a complex connected by an imine bond (between the aldehyde and the primary amine) and two boronate ester bonds (between the boronic acid and the diol). The two resulting five-membered rings 48  should thus be connected via a central boron atom.  Scheme 4.5 Derivatization of 1,2-diols with the boronic acid-based three-reagent system.  If the stereoconfiguration of the diol was changed, the chemical shifts of the protons adjacent to the two five-membered rings were most likely to be affected. In the products of this reaction, candidate protons included the three α-protons of the diol (B), the imine proton (A) and the methyl protons on the amine (C) (Figure 4.4). In the NMR spectrum of 4-8, two peaks of equivalent abundances were observed for proton A (Figure 4.4), which should be a singlet prior to derivatization, suggesting that the two isomers were distinguishable using this method. It should be noted that excess amount of derivatization reagents were found to give rise to peaks overlapping with the signals of proton A in NMR, rendering the assignment of absolute configuration extremely hard. Attempts to separate the product from excess amount of reagents by TLC failed, as no product could be recovered from silica gel. This result was probably due covalent bonding of the reagent 49  with the TLC matrix, as silica hydroxyl groups may form boronate ester bond(s) with the boron atoms and therefore reverted the product to starting material.   Figure 4.4 NMR spectrum (300 MHz) of 4-8.  The two enantiomerically enriched diols 4-1 and 4-2 were then derivatized (Scheme 4.5) and analyzed using the same method. No differences in either the splitting patterns or the chemical shifts were observed for the NMR signals of the diol α-protons (proton B) and the methyl protons of the chiral amine (proton C) (data not shown). However, the higher-field peak of the two imine proton signals detected for the racemic sample was greatly decreased for 4-9, and in contrast to that, for 4-10, the lower-field peak was significantly reduced (Figure 4.5). Therefore, the stereoconfigurations could be assigned based on the signal of the imine proton. The Δδ of 0.047 ppm measured here was similar to the values reported in literature.84b Since the two peaks were well separated from each other, the enantiomeric excess (ee) of each diol could be calculated from peak area integrations. For 4-1 and 4-2, the ee values were determined to be 50% and 66%, respectively.    50   Structures -N=C(H)- 8.15 – 8.35 ppm 4-8 4-9 4-10 Derivatized 1,2-diols from cosmos petal wax Figure 4.5 Signature 1 H-NMR (400 MHz) signals of the 1,2-diols derivatized using boronic acid based three-reagent system.  The exact amount of 1,2-diols used in the reaction was determined by GC-FID using C24 alkane as inner standard. Derivatizing reagent 4-6 and 4-7 in the right stoichiometry were added to avoid problems with NMR signal interferences from excess reagents. The resulting sample was analyzed as described above. The higher-field peak of the two imine proton signals was clearly more abundant, almost identical to the peak pattern of 4-10 (R-diol standard) (Figure 4.5). This result for the stereoconfiguration of the petal wax diols thus qualitatively matched that obtained using Mosher’s method. The ratio of the two peaks was 1:4.3, indicating approximately 81% of all the 1, 2-diols possessed the R configuration, and only 19% had S configuration (ee = 62%), comparable to the percentage obtained 51  using the Mosher’s acid method.  4.3 Discussion 4.3.1 Comparison of the two derivatization methods The two NMR-based methods used here both proved capable of distinguishing the two diol stereo-isomers and gave consistent results. Both methods rely on chiral derivatization agents (CDAs), the aryl substituents in which provide an anisotropic magnetic effect to differentiate sterically inequivalent protons.  Mosher’s acid and acid chloride are probably the most commonly used CDAs, with intensive configuration studies carried out to understand and predict experimental results. For mono-functional compounds, stereoconfigurations can be determined without comparing to enantiomerically pure standards.76 For bifunctional compounds, relatively less is known about the configuration of their Mosher esters and therefore in most cases standards are necessary to assign absolute stereoconfigurations. As illustrated in this experiment, Mosher’s acid chloride reacts mostly with primary rather than secondary alcohol groups. Hence, harsher conditions are required to achieve bis-derivatization. Even though bis-MTPA ester derivatives have been reported for 1,2-diols,92 examples of using mono-MTPA ester derivatives to study the stereoconfiguration of adjacent hydroxyl93 or methyl groups94 are also found in the literature. The introduction of only one MTPA group might also provide the advantage of simplifying the splitting patterns in NMR spectra.  Although the signals of excess amounts of reagents do not typically overlap with those of diol α-protons in NMR spectra, the Mosher ester products were usually purified by flash chromatography or TLC prior to NMR analyses, as illustrated by various literature reports.76 In the experiments presented here, the protons of the two purified isomers were well separated in the spectrum and could be clearly identified; however, peak splitting into multiplets caused overlap. This problem can be overcome by switching to other 52  Mosher-like CDAs, such as AMA 80, which result in better separated proton signals.  Boronic acid-containing reagents have long been used to derivatize diols.81a One of the major advantages of this method is the formation of five-membered dioxaborolane rings, circumventing the kinetic resolution problems caused by the competition between the of two alcohol groups when forming monoesters.84b The assembly reaction only takes 10-25 min to complete, so the whole experiment can be finished within an hour.85 As shown by the current experiments, for aliphatic 1,2-diols, only the imine protons can be differentiated by NMR. However, for more substituted diols, multiple sets of protons adjacent to the five-membered rings can further be used to assign the two isomers.84b One problem of this method encountered during the analysis is that the reaction stoichiometry needs to be carefully calculated, because excess amounts of derivatization reagents will cause peak overlapping with the imine protons and purification is difficult. That requires the amount of diols be accurately measured, especially in the case of natural products isolated in small quantities from biological sources. Another disadvantage of this method is that little is known about the configuration of the assembly products, making the stereoconfiguration assignment rely entirely on synthetic enantiomer standards.  One disadvantage shared by the two NMR-based methods is that they can only afford an averaged result of the 1,2-diol mixture and hence do not provide the information of each homologue. In order to resolve the homologues, alternative GC-based method need to be considered instead. The enantiomers of different chain lengths could be separated on chiral capillary columns, and the enantiomeric and racemic standards generated in the present work can serve as reference for such future chromatographic work.  4.3.2 Possible biosynthetic pathway to 1,2-diols in cosmos The biosynthetic pathways to long-chain and very-long-chain1,2-diols are currently unknown. It is conceivable that the hydroxyl groups are introduced consecutively through 53  α-oxidation of either a primary or a secondary alcohol. However, to the best of my knowledge, such biosynthetic reactions have not been reported so far. Two more pathways are conceivable, one introducing both functional groups simultaneously, and the other one modifying pre-existing 1,2-functionalities. Since the evidence provided here on the stereoconfigurations of cosmos petal 1,2-diols strongly favors the latter two pathway alternatives, they will be discussed in more detail. Both pathways were different from those discussed in section 1.4., and had never been used to explain the biosynthesis of wax compounds.  The first possibility relies on oxidation of 1-alkenes into 1,2-epoxides followed by hydrolysis (Figure 4.6).95 Enzymes have been discovered from bacteria and yeast that can selectively hydrolyze short-chain, unbranched aliphatic epoxides ranging from C4 to C8 to yield R-1,2-diols.96 Upon feeding with racemic epoxides only R-epoxides were transformed, with retention of configuration in the hydrolysis step.96 The ee values for the unreacted S-epoxides were determined to be over 98%, and those for the R-diol hydrolysis products were between 35%-86%,96 similar to the diols from C. bipinnatus petal wax. Similar enzymes were found in bacteria and fungi that could selectively hydrolyze S-2-methyl-1,2-epoxides, and the products were characterized to be S-2-methyl-1,2-diols.97 In plants, although epoxide hydrolases have not been reported, the existence of similar enzymes is not unlikely.  Besides that, in 1972 and 1978, Kolattukudy reported evidence for another pathway leading to 1,2-diols. Long-chain α-hydroxy fatty acids (or their CoA esters) were reduced to corresponding 1,2-diols by an enzyme found in Zonotrichia leucophrys (white-crowned sparrow) (Figure 4.6).98 Even though further characterization of the enzyme has not been reported afterwards, the original work pointed to another possible biosynthesis pathway for 1,2-diols. This precursor-product relationship is further supported by the R-configurations of 1,2-diols discovered in the current project, as all of the unbranched hydroxy fatty acids resulting from fatty acid α-hydroxylation also possess R-configurations.99 54   Fatty acid α-hydroxylation occurs widely in both sphingolipid biosynthesis and fatty acid degradation. The hydroxylation reaction has been studied using cell extracts of mammals, plants and algae.99 A broad substrate spectrum was found for this type of reaction, ranging from short-chain99e via long-chain99a, 99c, d to very-long-chain fatty acids (or their CoAs).99b For unbranched fatty acids (CoAs), the products were always enantiomerically pure R-hydroxy fatty acids. However for the 3-methyl fatty acid (CoA) intermediate in the fatty acid degradation pathway, the position of the hydroxyl group was determined by the stereoconfiguration of the methyl group on the neighboring β-carbon.100 When it had the R configuration, only the 2S,3R product was observed, and vice versa.  Fatty acid 2-hydroxylase (FA2H) from sphingolipid biosynthesis was recently identified from mammalian cells.101 In vivo assays with FA2H overexpressing cell lines gave exclusively R-2-hydroxy palmitic acid,102 similar to the reaction carried out by cell extracts. Two similar genes AtFAH1 and AtFAH2 in the model plant Arabidopsis were characterized to have fatty acid 2-hydroxylase activity, and were thought to be involved in sphingolipid biosynthesis.103 The substrates of AtFAH1 were mainly very-long-chain fatty acids, whereas AtFAH2 only reacted with palmitic acid.103 However, stereochemical information is still missing for the two plant gene products.  Between the two possible pathways that lead to 1,2-diols, the second one seems more likely, because no epoxide hydrolase has been found in plants, and the epoxide hydrolase from other organisms characterized to date usually prefer short chain lengths. However, for the second pathway, although some evidence has been reported and also VLC 2-hydroxy fatty acids have been discovered, key information about the enzymes is missing as well. Hence, more biochemical evidence is still needed to prove the second pathway.  55   Figure 4.6 Hypothetical biosynthesis pathways to 1,2-diols and 1,2-diol-monoacetates. 1,2-Diols can be biosynthesized by epoxidation of alkenes followed by hydrolysis, or by α-hydroxylation of fatty acids followed by reduction of the acid group into hydroxyl group. 1,2-Diol-monoacetates can be biosynthesized by esterification directly from 1,2-diols, or by esterificaion of the α-hydroxy acid, followed by reduction.  4.3.3 Possible biosynthetic pathway to 1,2-diol monoacetates in cosmos The chain length profiles of monoacetates and free 1,2-diols were very similar, suggesting that the two compound classes were biosynthetically linked. The biosynthesis of 1,2-diol 1-acetates could proceed directly from 1,2-diols with the help of an acyltransferase (Figure 4.6). The same enzyme may also be responsible for the formation of 1,2-diol 2-acetates, or alternatively, a second acyltransferase with high regioselectivity towards secondary alcohol groups in the presence of primary alcohol is involved in the biosynthetic pathway. Such an enzyme is not unprecedented. A series of acyltransferases (GPATs) were identified in Arabidopsis that catalyze the acylation of glycerol-3-phosphate (G3P). Three enzymes in this family, GPAT-4, GPAT-5 and GPAT-6, when assayed with α,ω-dicarboxylic acid CoA esters and [14C]-G3P, gave products acylated in the sn-2 position of G3P exclusively.104 Further studies indicated that GPAT-4 and GPAT-6 had strong substrate preference 56  towards C16:0 and C18:1 ω-oxidized acyl-CoAs105, and they are therefore thought to be involved in cutin biosynthesis.104-105 In contrast, GPAT-5 was hypothesized to be associated with suberin biosynthesis,104-105 and its substrates varied more in chain lengths.105 Possibly a similar enzyme, present in Cosmos bipinnatus petals, is able to preferentially transfer an acetyl group onto the secondary hydroxyl group of 1,2-diol substrates.  Another explanation for the biosynthesis of 1,2-diol 2-acetates involves the hypothetical precursor of 1,2-diol (Figure 4.6). If α-hydroxy fatty acids (or CoAs) are indeed the precursors of 1,2-diols, then an acyltransferase might first convert the α-hydroxy fatty acids (CoA) into α-acetate fatty acids (CoA), and then a fatty acyl reductase (FAR) would reduce the acyl group into the primary hydroxyl functionality. The same reductase might also catalyze the reduction of α-hydroxy fatty acids (CoA) to diols. Currently evidence is lacking for both hypothetical pathways. Therefore, further investigations are required to search for possible pathway intermediates and to identify key enzymes.            57  5 β-Keto acid derivatives: potential pathway intermediates and end products 5.1 Introduction Presence of β-functionalized wax compounds in plants The work presented in chapters 3 and 4 focused on 1,2-bifunctional wax compounds. Besides them, wax constituents with β-functionality are also naturally present and are categorized as primary/secondary bifunctional compounds (section 1.4.3). However, wax compounds with functional groups in the β position were only found in very few plant species so far. All of the identified β-functionalized wax compounds have predominantly even-numbered chain lengths, and the majority of them fall into the compound class of 1,3-alkanediols. For example, 1,3-diols with chain lengths ranging from C20 to C28 were found in the leaf wax of Ricinus communis,106 the petal wax of Cosmos bipinnatus (see Chapter 3)52 and the leaf wax of Papaver alpinum.107 Besides 1,3-diols, the only other 1,3-functionalized compound class identified so far are the β-hydroxy aldehydes from the leaf wax of Ricinus communis.106 A systematic search of 1,3-functionalized compounds in plant cuticles is necessary to reveal the full compound class spectrum of cuticular waxes.  The 1,3-configuration of functional groups common to some compound classes suggests that they are biosynthetically related. It seems plausible that they are formed by modification of the 1,3-functionalized intermediates occurring during common wax biosynthesis, namely the -ketoacyl-CoA, -hydroxyacyl-CoA or 2,3-enoyl-CoA esters formed during fatty acid elongation (Figure 5.1). The biosynthesis of 1,3-diols would thus require only one or two enzymes, one of them reducing the head group to an alcohol and the other one generating the -hydroxy group if it is not present yet. The primary functional group can be reduced by an enzyme similar to CER4, and the β-carbonyl group can be reduced by KCR-like enzyme to generate secondary alcohol. This is just one example of pathway hypothesis. Alternative pathways that use other 1,3-functionalized intermediates 58  are also possible, for example, via β-keto or β-hydroxy aldehydes. The essential feature of all these hypothetical pathways is that intermediates must escape from the fatty elongation cycle, or they must be intercepted by other enzymes. However, it is generally assumed that the KCS, KCR, HCD and ECR enzymes of the elongase are tightly associated and substrate is channeled between them.  Figure 5.1 Hypothetic alkane and 1,3-alkanediol biosynthesis pathways via β-ketoacyl-CoA intermediates. The β-keto acid can be derived from the elongation cycle and undergo instantaneous decarboxylation to afford alkan-2-one. The alkan-2-one can be reduced to alkane by a mechanism similar to the elongase reduction sequence. The reduction of β-ketoacyl-CoA to 1,3-diol can be achived by two enzymes: CER4-like and KCR-like reductases. Alternatively, alkane and 1,3-alkanediol can also be biosynthesized via fatty acid acyl-CoA intermediates (colored in grey).  It is noteworthy that β-functionalized wax compounds may also serve as intermediates in the biosynthesis of further wax constituents including alkanes. The existence of such a secondary alkane pathway is suggested by the finding that Arabidopsis cer1cer3 double 59  mutants do not show complete depletion of alkanes.108 In such an alternative alkane pathway, a functional group at the C-3 position may assist the C-C bond cleavage. For example, the β-ketoacyl-CoAs elongation intermediates can lose the terminal carbon via decarboxylation, and give rise to odd-numbered alkan-2-ones (Figure 5.1). The transformation of the resulting ketones to the corresponding alkanes might be achieved by a sequence of reductions similar to the elongation cycle. Admittedly, without the assistant of the adjacent carbonyl groups, the last two reduction steps, dehydration of hydroxyl group and reduction of alkene, are very difficult reactions for enzymes. However, this hypothesis cannot be ruled out at this point due to recent discovery of possible pathway intermediates, alkane-2-ones and alkane-2-ols, in the potato leaf wax.109  To test the above pathway ideas, a first step is to identify possible pathway intermediates. However, there are no available techniques that we can use to detect the intermediates from the internal lipids of plants, where the biosynthesis of wax compounds occurs. As the fallback alternative, wax analysis of plant cuticles may provide some clues, because chances are that some pathway intermediates escape to the cuticle. To find and identify these intermediates in cuticular waxes, standard compounds are required. In addition to that, the search of β-functionalized wax compounds in plant waxes can also be greatly facilitated if standard compounds are produced.  The bifunctional VLC substrates are not commercially available, and therefore the only way to provide the standard compounds is by chemical synthesis. However, even though the synthetic tools for similar reactions have been published for decades, their applications to plant wax studies have only been scarcely reported. To the best of my knowledge, the only very-long-chain β-functionalized standards synthesized were β-ketotetracosanoic acid ethyl ester, tetracosane-1,3-diol and β-hydroxytetracosanal,106 which were only a small portion of all possible structures. Therefore, in order to identify other β-functionalized wax compounds such as β-keto acid derivatives from plant waxes, more synthetic substrate chain lengths and functional group combinations are needed. 60   5.2 Results Work in this project proceeded in two steps: Initially, methods and conditions for synthesizing long-chain and very-long-chain β-functionalized standards were developed (section 5.2.1). The synthesized standards were then used to identify the structures of unknown compounds found in aloe waxes (5.2.2).  5.2.1 Synthesis of β-functionalized wax compounds Wax compounds are derivatives of very-long-chain fatty acids (VLCFAs), the chain length of which ranges from C24 to C34. Arabidopsis waxes are dominated by C30 compounds, and therefore this chain length was chosen for the target structures in this study. Due to the costs of C30 starting materials, the synthetic strategies and reaction conditions were first tested using long-chain starting materials (C18). The synthesized long-chain standards can also be used as model compounds to help better understand the fragmentation patterns in MS. The major challenge in these syntheses is posed by the low solubilities of the starting materials, which sometimes require optimization of solvents and reagents.  The method used to introduce the β-functional group into the substrates was first examined. Taking into account available synthetic methods and possible downstream products, β-keto ester was chosen as the target β-functionalized intermediate. Among various methods to synthesize β-keto esters,110 Meldrum’s acid method was relatively easy to carry out, and it was suitable for long-chain aliphatic substrates (C14 and C16 acids). 111 Stearic acid was activated using DCC/DMAP and coupled with Meldrum’s acid 5-2 to give the product 5-3 with 77% yield (Scheme 5.1).  61  Scheme 5.1 Synthesis of octadecanoyl-Meldrum’s acid 5-3 and its derivatives.  Methanolysis of the acyl-substituted Meldrum’s acid proceeded smoothly with the evolution of acetone and carbon dioxide. After 3-4 hours, all starting material was transformed into methyl 3-ketoeicosanoate 5-5, and the product could be purified via recrystallization (yield = 71%). If replacing methanol with other nucleophiles, a similar ring opening process could occur to afford different β-keto esters, thioesters, and amides.110, 112 N-acetyl cysteamine (NAC), was reacted with 5-3 under similar conditions to afford β-keto NAC-thioester 5-4 (yield = 79%) (Scheme 5.1).  The synthetic conditions to further modify the intermediate 5-5 by hydrolysis and reduction into other β-functionalized compounds were then studied (Scheme 5.2). For short-chain β-keto esters, hydrolysis had usually been carried out in basic conditions at ambient temperature.113 However, no hydrolysis product of 5-5 could be detected after 3 days using similar conditions, indicating reduced reactivity for long-chain substrates. Raising the reaction temperature could facilitate the hydrolysis, however it also led to the decarboxylation of the hydrolyzed product. When the reaction temperature exceeded 60°C, only the decarboxylation product could be detected. Alternative conditions for acid-catalyzed hydrolysis114 were then tested. The methyl ester 5-5 was dissolved in a mixture of acetic acid and HCl, and the hydrolyzed product 5-6 precipitated out gradually over time (yield = 51%). Reflux of 5-6 in acetone caused decarboxylation and afforded 62  methyl ketone 5-7 quantitatively. β-Hydroxy ester 5-8 was obtained by reducing 5-5 with sodium borohydride (yield = 76%). Its hydrolysis product 5-9 was synthesized using the same acid hydrolysis conditions as for 5-6, giving a yield of 58%.   Scheme 5.2 Synthesis of long-chain β-functionalized compounds from methyl 3-oxoeicosanoate 5-5.  The above synthetic conditions were then applied to the synthesis of VLC β-functionalized compounds. Although some conditions were still compatible, many conditions needed to be adjusted or changed due to the reduced solubilities (and therefore reactivities) of substrates. When the same conditions (DCC/DMAP) were used in the coupling reaction of octacosanoic acid 5-10 and Meldrum’s acid, considerable amounts of side-products were observed. This problem could be circumvented if the coupling reagent DCC was replaced by EDC, and the octacosanoyl Meldrum’s acid 5-11 was obtained with excellent yield (91%) (Scheme 5.3). The ensuing methanolysis step was performed under the same conditions as for long-chain substrate, except that a longer reaction time (8 h instead of 3 h) was required.   Scheme 5.3 Synthesis of octacosanoyl-Meldrum’s acid 5-11 and methyl 3-oxotriacontanoate 5-12.  63  The acid hydrolysis method had to be abandoned since 5-12 had very low solubility in acetic acid, even under elevated temperature. For basic hydrolysis, no reaction was observed if the reaction was kept at 66°C (THF). However, once the temperature was raised to 120°C, complete hydrolysis could be achieved within hours (Scheme 5.4). The resulting β-keto acid underwent decarboxylation simultaneously to give the methyl ketone 5-13 with 73% yield. Subsequent NaBH4 reduction afforded 2-nonacosanol 5-14. VLC β-hydroxy ester 5-15 was synthesized similarly to 5-8, with a yield of 72%. The hydrolysis of 5-15 was also tested using similar basic hydrolysis, and the product 5-16 could be identified by comparing the mass spectrum to that of 5-8. The yield was approximately 85% based on the GC-MS peak integrations. However, very little product could be isolated by TLC purification. Optimization of the purification method is currently underway. The mass spectra of all C30 standard compounds are summarized below (Figure 5.2).    Scheme 5.4 Synthesis of very-long-chain β-functionalized compounds from methyl oxotriacontanoate 5-12.       64      Figure 5.2 Mass spectra of the TMSi ethers of C30 standard compounds .  5.2.2 Identification of β-functionalized wax compounds in Aloe arborescens leaf waxes In the on-going project of Aloe arborescens wax analyses, a series of unknown compounds were detected by GC-MS. The unknown constituents were present in considerable amounts in the waxes, and consisted of three homologous compounds (Figure 5.3). Under 5-12 5-13 5-14 5-15 65  total ion monitoring, only the second and the third homologues could be detected, as the first homologue co-eluded with the most abundant wax constituent, C31 alkane. However, it could be visualized using the selective ion monitoring (SIM) mode, monitoring the signature m/z 175 ion (Figure 5.3). The mass spectra of the TMSi derivatives of the unknown compounds showed fragment ion m/z 73 [OTMSi]+, and no obvious ion m/z 103 [CH2OTMSi] +, implying a possible secondary alcohol functionality (Figure 5.4). The fragment ion m/z 175 was of high intensity in the mass spectra of all three homologues. Another characteristic of the mass spectra were chain length specific base peaks separated by 24 mass units: m/z 491, 515, 539, respectively, and daughter ions with 58 mass units less than the base peaks.   Figure 5.3 GC-MS chromotograms of the TMSi ethers of Aloe waxes. (A) Full scan mode, (B) SIM mode monitoring fragment m/z 175.  A B 66   Figure 5.4 Mass spectra of unknown compounds from A. arborescens leaf waxes. (A) TMSi ether of the first unknown homologue. (B) TMSi ether of the second unknown homologue. (C) TMSi ether of the third unknown homologue.  Based on the mass spectra of the unknown compounds, it seemed plausible that they were β-hydroxy fatty acid methyl esters. This may be illustrated using the mass spectrum of the third unknown homologue as an example (Figure 5.4, C): The base fragment m/z 539 [M-15]+ may be interpreted as loss of CH3 from the TMSi-derivatized alcohol group. Its daughter ion m/z 481 [C27H55CH(OTMSi)] + could be explained as the α-fragment cleaved between C-2 and C-3 (Scheme 5.5). The signature fragment ion m/z 175 [CH(OTMSi)CH2COOMe] could then also arise from α-fragmentation, but between C-3 and C-4. This proposed fragmentation pattern suggested an overall structure of 3-hydroxytriacontanoic acid methyl ester. The mass spectral assignments are analogous to A B C 67  those of reported wax compounds with similar structures, such as 5-hydroxyaldehydes.47 To test this hypothesis, the mass spectrum of standard 5-15 was compared to that of the third unknown homologue, and they were found identical. A co-elution experiment further confirmed that the third homologue in the unknown series was methyl 3-hydroxytriacontanoate, as the retention times of both compounds on the GC-FID were identical (Figure 5.5).  Scheme 5.5 Fragmentation patterns of methyl 3-hydroxytriacontanoate.  Figure 5.5 Co-elution expermient of the A. arborescens leaf wax and β-hydroxy acid methyl ester standard 5-15. The retention times of the third unknown homologue and 5-15 were approximately the same on GC-FID.  To exclude other alternatives, the synthetic precursor of 5-15, β-keto methyl ester 5-12, A. arborescens leaf wax 5-15 68  was also included in the co-elution experiment. The retention time of 5-12 was very close to the peak right after methyl 3-hydroxytriacontanoate (data not shown). However, their mass spectra shared no similarity. In addition, a search of signature fragment of β-keto methyl ester, the m/z 173 ion [C(OTMSi)CHCOOMe]+, using the SIM mode showed no significant signals in the GC-MS chromatogram of the aloe wax. Hence, it could be concluded that β-keto acid methyl esters were not present in aloe wax. Further attempts were made to search for other hydroxy acid methyl ester isomers in the aloe wax using the SIM mode. In the mass spectrum of 5-15, cleavage between the hydroxyl-substituted carbon and the adjacent carbon resulted in a signature α-fragment, i.e., m/z 175. Similarly, the 2-, 4-, 5-, 6- and 7-hydroxy acid methyl esters should also have α-fragments m/z 161, 189, 203, 217 and 231, respectively. However, no significant signals could be detected for any of these fragment ions. Hence, it could be concluded that there were no positional isomers of β-hydroxy acid methyl esters in the aloe waxes.  Beside the identified methyl 3-hydroxytriacontanoate homologue, the other two unknown homologues could be identified as methyl 3-hydroxyoctacosanoate and methyl 3-hydroxyhexacosanoate, based on their base peaks (figure 5.4, A, B). According to the GC-MS-SIM chromatogram for m/z 175, the most abundant homologue was methyl 3-hydroxyoctacosanoate, and the C27 and C31 homologues were roughly 13% and 20% of its abundance, respectively. Further quantifications are currently underway (pers. communication A. Luna and R. Racovita)  5.3 Discussion 5.3.1 Synthesis of β-functionalized wax compounds A series of β-functionalized long-chain and very-long-chain wax compounds were synthesized with acceptable yields. The same synthetic strategy was applied to both the C20 and C30 products. However, due to the change in the substrate chain lengths, two sets of different reaction conditions were developed. For future synthesis of the substrates with 69  chain lengths between C20 and C30, at least one of the reaction conditions should give satisfactory result.  Central to this synthetic strategy is the Meldrum’s acid method. A similar method using malonic ester had been used previously to synthesize β-ketotetracosanoic acid ethyl ester.106 Both methods have broad chain length compatibility, however the advantage of the Meldrum’s acid method is that the acyl-derivatized Meldrum’s acid can react with various nucleophiles. Among different nucleophiles, NAC is of particular value to future researches, as it can serve as a surrogate to coenzyme A. Examples had been reported that some polyketide biosynthesis enzymes were able to accept and process NAC derivatives, as substitutes for CoA derivatives.115 The β-functionalized acyl NACs obtained from the Meldrum’s acid method can therefore be used in future biochemical assays to test pathway hypotheses or to study the biosynthesis of β-functionalized wax compounds.  Furthermore, the obtained β-functionalized compounds can serve as standards to help the identification of pathway end products and intermediates from plant waxes, which was the main purpose of this study. One illustration of that is the identification of the unknown compounds found in the aloe waxes. The waxes from other plant species can be analyzed in a similar way. In fact, a project into potential β-functionalized compounds in Arabidopsis waxes is currently underway, as these might occur as intermediates on an alternative alkane biosynthesis pathway: alkane-2-ones and alkane-2-ols (Figure 5.1). The β-functionalized compounds can also be used as synthetic intermediates to produce enzyme substrates for in vivo assays (see Chapter 6).  5.3.2 Biosynthesis of β-hydroxy acid methyl esters in A. arborescens β-Hydroxy acid methyl esters had not been previously found in plant waxes, but methyl esters are known to occur.49a, 116 Methyl esters could be biosynthesized by acyltransferases,117 or more likely by methyltransferases, as illustrated by the 70  methyl-esterification of homogalacturonan.118 In this example, S-adenosyl-methionine (SAM) served as the donor of methyl group. On the other hand, nothing is known about the biosynthesis of the β-hydroxy group. However, based on the current knowledge about the biosynthesis of bi- or multifunctional wax compounds, it can be hypothesized that the β-hydroxy functional group is introduced either during the elongation cycle, or alternatively by oxidation of the β-methylene in the wax compound that has primary functionality (Figure 5.6).   Figure 5.6 Hypothetical biosynthesis pathways to β-hydroxy acid methyl esters. The methyl ester can be synthesized by an methyltransferase or acyltransferase from the corresponding acyl-CoA. The β-functional group can be introduced from the elongation or be installed via hydroxylation.  There are few possible hypothetical reaction sequences. If the KCS product β-ketoacyl-CoA is exported from the elongation cycle, then the keto group may be reduced 71  to produce the β-hydroxy functionality. It is also possible that the β-hydroxyacyl-CoA intermediate of elongation is used directly to synthesize the β-hydroxy acid methyl ester. Comparing the two possibilities, the first one seems less likely, because the intermediate β-keto acid methyl esters were not found in the aloe wax. As for the third hypothesis, the β-hydroxy acid methyl esters can be directly synthesized from VLC acyl-CoAs by β-hydroxylation. It can also be produced on a pathway similar to fatty acid β-oxidation, where the fatty acyl-CoA is first transformed into 2,3-enoyl-CoA, and then the 2,3-enoyl-CoA is hydrated to afford 3-hydroxyacyl-CoA.119 To further test these pathway hypotheses, intermediate compounds need to be identified and also the related genes be cloned and characterized.                   72  6 Synthesis of isotope-labeled very-long-chain aliphatic substrates and preliminary in vivo assays of CER1 in yeast (Saccharomyces cerevisiae) 6.1 Introduction Biochemical in vivo or in vitro assays are currently the most powerful tools to determine the function of certain enzymes. Assays with substrate feeding are superior to those relying on endogenous substrate availability. For the wax biosynthesis enzymes, substrate feeding assays are difficult to conduct in vitro, mainly because the involved enzymes are membrane-bound. It is generally assumed that the highly hydrophobic substrates must be delivered directly from a membrane environment to the enzyme active site, which is difficult to mimic using in vitro assays with solubilized enzymes. The most common solution is to express the enzymes in a heterologous system, like yeast and bacteria, and conduct the assays in vivo by supplementing substrate exogenously. Several wax-related enzymes were successfully studied using this approach, such as WSD1 in wax ester biosynthesis,27 KCS in VLCFA elongation,12, 120 and FARs in VLC primary alcohol biosynthesis.121  CER1 plays a pivotal role in Arabidopsis alkane biosynthesis, and is thought to be the decarbonylase converting aldehydes into the final products (see section 1.3.2). In vivo assays of CER1 and CER3 in yeast confirmed that the two enzymes are required for the alkane biosynthesis pathway. However, information regarding their substrates is still missing, and hence final evidence from in vivo feeding experiments using heterologously expressed CER1 or CER3 is needed.  For feeding experiments, all alternative substrates must be prepared. However, the sources of VLC substrates are very limited. In a few cases, substrates (mainly VLC fatty acyl-CoAs) can be provided by the heterologous system.33c, 121 Commercially available VLC compounds are mainly the fatty acids and alcohols, which make up only a small percentage of all possible substrates required by wax biosynthesis enzymes. VLC fatty aldehydes, for example, are not commercially available, and have to be prepared by 73  chemical synthesis.  Isotope labeling is very useful in tracing the enzymatic product of interest, and can even provide evidence for possible reaction mechanism. However, the VLC isotope-labeled substrates are even less accessible. Many enzymes involved in Arabidopsis wax biosynthesis (likely) prefer C30 substrates, as evidenced by the phenotypes of various mutants.21, 122 For CER1 and CER3, the drastic increase of C30 aldehyde and C30 alcohol in the cer1 and cer3 mutants, respectively, strongly suggests that C30 is the preferred substrate chain length for the enzymes.34 Yet, to the best of my knowledge, no isotope-labeled C30 aliphatic substrates have been used to study wax biosynthesis.  The main goal of this project was to provide C30 isotope-labeled VLCFA derivatives, which can serve as substrates for future assays. Cold isotope labeling was chosen over radioactive isotope labeling because non-radioactive isotopes (2H, 13C, 18O) can be handled in normal environments, and because they can be easily detected by readily available mass spectrometric detection techniques using higher mass isotopic peaks. The carbon in β position to the VLC substrates’ primary functionality is an ideal site where isotope like deuterium (2H) can be introduced, as (1) methods for introducing β-functional groups into wax compounds has been successfully established in Chapter 5, and (2) the β protons are not particularly acidic and the isotope label(s) are therefore unlikely to be lost during the enzyme reaction. As a further goal, the unlabeled VLC fatty aldehyde substrate for CER1 also had to be synthesized to explore suitable in vivo assay conditions.  6.2 Results Work in this project proceeded in three steps, first with the synthesis of labeled substrate (section 6.2.1), then the preparation of the aldehyde substrates (section 6.2.2), and finally initial attempts at using the substrates in CER1 in vivo assays (section 6.2.3).  74  6.2.1 Synthesis of deuterium-labeled very-long-chain substrate The deuterium could be introduced using ketones or alcohols as starting materials. Ketones needed to be transformed into the corresponding tosylhydrazones, and then reduced by deuterated reagents like NaBD4 or LiAlD4. 123 Alcohols also needed to be first converted into good leaving groups, such as tosyl and iodo groups, followed by reduction with NaBD4 or LiAlD4. 124 Based on available β-functionalized substrates, possible synthetic strategies were considered (Scheme 6.1). The first strategy required to first synthesize the β-tosylhydrazone acid methyl ester from β-keto acid methyl ester. However, when β-keto methyl ester 5-5 was reacted with tosylhydrazide, the desired product was not formed, probably because of interference from the ester carbonyl.  Scheme 6.1 Retrosynthetic analysis of β-D-labeled substrate. Next, the method to transform the β-hydroxyl group in 5-8 into iodine was tested (scheme 6.2). The β-hydroxyl group of 5-8 was reacted with triphenylphosphine, imidazole and iodine.125 The desired product 6-1 was indeed formed, based on the NMR spectrum of the crude product. However, a side product with very similar Rf value to the iodide product 6-1 was found on TLC, and it could not be separated. NMR and mass spectra of the crude product indicated the side-product was α, β-unsaturated ester 6-2, which was very likely generated by elimination of hydroiodic acid. Reducing the crude product with LAH exclusively afforded the elimination product 6-2. 75   Scheme 6.2 Isotope-labeling strategy via β-iodide intermediate. Only elimination product was obtained. For the tosylation of β-hydroxy acid methyl ester (Scheme 6.3), various reaction conditions (base, solvent, temperature) had been mentioned in the literature.126 The optimal condition that gave the best yield was using triethylamine (TEA) and DMAP at room temperature overnight. However, the conversion percentage was still relatively low (yield = 48%). The unreacted starting material 5-8 could be recycled. Addition of DMAP was necessary, as the yield was greatly decreased when using TEA alone. Switching to strong base like potassium tert-butoxide or raising reaction temperature gave rise to elimination product. Deuterium could be introduced by reducing the tosylated intermediate with NaBD4. 127  The transformation from 6-3 to 6-4 went smoothly with a yield of 83%. The overall yield of the two steps was 40%.  Scheme 6.3 Isotope-labeling strategy via β-tosyl intermediate. The β-position labeling was verified by NMR and GC-MS (Figure 6.1). In the NMR spectrum of 6-4, the signal of the α-methylene protons was split into a doublet, which in unlabeled compound should be a triplet, indicating that one of the β protons was replaced by deuterium. The isotope peaks of the molecular ions in the mass spectra of both unlabeled and labeled methyl esters were normalized to their M+ isotope peaks and 76  compared. The molecular ion of 6-4 was one mass unit higher than that of the unlabeled compound, confirming the introduction of one deuterium. However, small amounts of unlabeled compound (m/z 326) were also present in 6-4, with roughly 20% the intensity of m/z 327. No [M-1]+ peaks were found in both mass spectra. The intensities of [M+1]+ and [M+2]+ isotope peaks relative to the corresponding M+ peaks were roughly the same between two samples: 24% for [M+1]+ and 3.2% for [M+2]+. They both matched the calculated values. The percentage of labeled compound in 6-4 was calculated to be 83%, based on the intensities of m/z 326.2 and m/z 327.2 isotope peaks.   Figure 6.1 Charaterization of deuterium-labeled substrate 6-4. (A) 1 H-NMR spectrum (300 MHz) of 6-4; (B) The molecular ions of 6-4 (red bars) and the corresponding unlabeled ester (blue bars) in their mass spectra.  The same labeling strategy was applied to very-long-chain substrate (Scheme 6.4). The β-keto ester was prepared using Meldrum’s acid, as shown in Chapter 5. Reduction of 5-12 A B 77  with NaBD4 introduced a first deuterium atom, with a yield of 92%. 6-5 was turned into its tosyl derivative 6-6 using the same tosylation conditions as described above. Then reduction with NaBD4 incorporated a second deuterium, which increased the masses of isotopic fragments in the mass spectrum by one more mass unit. The yields for the last two steps were 64% and 58%, respectively. Attempts to optimize the reaction conditions were not successful. 100 mg of 6-7 were synthesized, with an overall yield from 5-12 of 34%. The purity of 6-7 was determined to be over 98% based on GC-MS.  Scheme 6.4 Synthesis of double-deuterium labeled very-long-chain substrate. The NMR spectra of 6-7 confirmed the introduction of two deuteriums at the β position, as no splitting was observed for the α-methylene protons (Figure 6.2, A). The small shoulder on the left of the α-methylene proton signal was assigned as the mono-labeled product, based on the chemical shift. The mass spectra of unlabeled and double-labeled C30 methyl ester were analyzed similarly as mentioned above (Figure 6.2, B). An increase in the mass of the molecular ion of 6-7 by two mass units further corroborated the double labeling with deuteriums. Based on the intensities of m/z 467.5 and 468.5 in the mass spectrum of 6-7, 83% double-labeled compound (m/z 468.5) was present, and the remaining 17% was mono-labeled compound (m/z 467.5). In both unlabeled and double-labeled compounds, the [M+1]+ and [M+2]+ isotope peaks were roughly 39% and 8% the intensities of their M+ peaks, respectively. The percentages were also very close to the calculated values. 78    Figure 6.2 Charaterizations of double-deuterium labeled substrate 6-7. (A) NMR spectrum (300 MHz) of 6-7; (B) molecular ions of 6-7 (red bars) and the corresponding unlabeled ester (blue bars) in their mass spectra.  6.2.2 Synthesis of very-long-chain fatty aldehyde VLC primary alcohols were commercially available. Therefore a one-step synthesis of the aliphatic aldehydes from the corresponding alcohols was possible.128 As previous syntheses of VLC substrates (see Chapter 5), the conditions were tested on long-chain substrate first. 1-Octadecanol 6-8, after stirring at room temperature with PCC for 1.5 – 2 h, was successfully oxidized into aldehyde 6-9, the structure of which was confirmed by both GC-MS and NMR. The best yield achieved was 67%.  A B 79   Scheme 6.5 Synthesis of long-chain and very-long-chain fatty aldehydes.  For the C30 substrate 1-triacontanol 6-10, if the same conditions were used, considerable amounts of 6-10 still existed after 8 h, probably due to low solubility of substrate in dichloromethane. Hence, the reaction temperature was elevated to reflux, under which conditions the substrate was completely converted after 2 h, as indicated by TLC. Reaction yield was over 90% based on the GC-MS chromatogram of the crude product. The isolated yield was only 47%, however further optimization of the purification step was not carried out. Ethanol-free chloroform can also be used as solvent in the reaction and gave comparable results.  6.2.3 Preliminary in vivo assays of CER1 using heterologous expression in yeast Preliminary in vivo assays of the function of CER1 were carried out using transgenic yeast (Saccharomyces cerevisiae) harboring the Arabidopsis CER1 gene. All yeast materials were provided by UBC Botany MSc student Mariya Skvortsova and the presence of the CER1 gene was verified by PCR (personal communication, Mariya Skvortsova). In each experiment, four parallel controls were included: (1) with C30 primary alcohol as substrate; (2) without C30 aldehyde substrate; (3) without CER1 gene (wild-type yeast); (4) without both. After 16 h of incubation, yeast lipids were extracted and analyzed by GC-FID and GC-MS after TMSi-derivatization. The GC chromatograms and mass spectra were analyzed and compared.  In one experiment, when C30 aldehyde was fed to CER1, C30 acid methyl ester was detected. The same compound was not found in all four negative controls. However, this result could not be repeated in three other independently conducted experiments. 80  Therefore, it was concluded that the detected C30 acid methyl ester was an artifact generated during lipid extraction or from contamination. In addition, the expected product of CER1, C29 alkane, was not found in any of the experiments (Figure 6.3). Unreacted aldehyde could be recovered from the medium supernatant, albeit in vastly varying percentages of the original amounts.  81   Figure 6.3 Co-elution expermient of the preliminary in vivo assays of CER1 in yeast. GC-FID chromatograms of assays with (A) C30 aldeyde fed to yeast expressing CER1; (B) C30 primary alcohol fed to yeast expressing CER1; (C) yeast expressing CER1 without substrate feeding; (D) wild-type yeast alone. (E) C29 alkane standard.  A B C D E 82  6.3 Discussion 6.3.1 Synthesis of double-deuterium labeled VLC substrates Two deuterium labels were successfully introduced in the β-position of VLC fatty methyl ester, as shown by NMR and GC-MS results. In this project, only the C30 substrates were synthesized. However, the developed labeling method can be easily applied to substrates with other chain lengths. The obtained double-labeled compounds can be converted into VLC fatty aldehyde as well as other VLCFA derivatives, which can serve as substrates for other wax biosynthesis enzymes.  In 6-10, roughly 83% of the methyl ester was characterized to be double-labeled product, and the remaining 17% was mono-labeled product. This raises the question whether the deuterium label in the mono-labeled product was introduced during the first NaBD4 reduction (from 5-12 to 6-5) or during the second reduction (from 6-6 to 6-7). To answer this question, the mass spectra of 6-5 and the unlabeled β-hydroxy acid methyl ester 5-15 were analyzed using the same method as above (Figure 6.4). The [M-15]+ ion of 6-5 was one mass unit higher than that of the unlabeled compound, corresponding to the mono-deuterium label. Only very small amounts of unlabeled compound (m/z 539.6) could be found in 6-5, suggesting the introduction of label was nearly complete. Therefore, it can be concluded that the deuterium of the mono-labeled compound in 6-7 originated from 6-5. This also means that the labeling efficiency from 6-6 to 6-7 was only 83%. Further investigation is required to understand the reaction mechanism.  83   Figure 6.4 The molecular ions of 6-5 (red bar) and 5-15 (blue bar) in their mass spectra.  6.3.2 Preliminary in vivo assays of CER1 in yeast The in vivo assays carried out in this study did not yield any enzymatic turnover product. The exact reasons for these negative results are currently unclear. In fact, similar feeding experiments were mentioned in the literature, and no enzymatic product was found as well.33b One explanation could be that the yeast cells failed to import the aldehyde substrate from the culture medium, and therefore the substrate was inaccessible to CER1 enzyme. This explanation was supported by the recovery of substantial amounts of C30 aldehyde from the culture supernatant. It should be noted that, even though the same feeding method had been successfully used for C24 and C28 primary alcohols, 27 it is still possible that the increased hydrophobicity of C30 aldehyde, compared to C28 alcohol, hindered substrate uptake. In future experiments, in vitro assays of yeast microsomal preparations or of protein extracts can be used to increase the availability of substrate to CER1 enzyme.  Another possible explanation is provided by Bernard et al.33c When CER3 alone was expressed in yeast, no aldehyde intermediate could be detected. Therefore, the authors proposed a biochemical model where CER3 and CER1 work synergistically in the production of alkane,33c with direct delivery of the aldehyde intermediate from CER3 to CER1. This hypothesis was partially supported by the observed physical interactions 84  between CER1 and CER3 enzymes in both yeast and plants.33c In the insect alkane biosynthesis pathway, similar enzyme associations have also been proposed.32 The P450 oxidative decarbonylase and the NADPH-cytochrome P450 reductase are thought to physically associate with fatty-acyl-CoA reductases, for the purpose of substrates channeling. To prove this hypothesis using the yeast system, two parallel experiments can be designed: either C30 aldehyde substrate is fed to the yeast strain that expressing both CER3 and CER1, or to a strain expressing only CER3. If C29 alkane is found only in the former experiment, then the function of CER3 as essential for delivering this substrate is confirmed.  Lastly, it is also still possible that CER1 is not functioning as a decarbonylase, i.e. its substrates are not VLC aldehydes. The evidence for the aldehyde decarbonylase function of this enzyme mainly came from mutant phenotype analysis, where the C30 aldehyde increased in cer1 mutants. It is still possible that plant alkanes are biosynthesized via some other pathway(s), for example, decarboxylation of β-functionalized compounds (see section 5.1). Further investigations are required to distinguish between the various pathway hypotheses.           85  7 Summary and future directions The plant cuticle is of great biological and ecological importance, as it serves as the first protection barrier of plants and the interface for plant-environment interactions. To fully understand its functions, the plant cuticle needs to be studied at a molecular level, with special focus on the wax compounds. As reviewed in Chapter 1, significant progress has been made recently in elucidating the biosynthetic pathways leading to wax compounds, especially those with primary functionalities. In contrast to that, current knowledge about multifunctional wax compounds is still very limited. Hence, the major objectives of this study were to expand the knowledge of the multifunctional wax compounds, particularly in 1,2- and 1,3-bifunctional compounds (see 1.5).  Through the research work presented in the previous chapters, contributions into the chemical composition and biosynthesis of bifunctional wax components have been made. Specifically, I have identified novel 1,2- and 1,3-bifunctional compounds from C. bipinnatus petal wax and A. arborescens leaf wax, respectively, based on synthetic standard compounds, and established two independent NMR-based approaches to analyze the stereoconfigurations of 1,2-alkanediols found in C. bipinnatus petal wax.  Besides, a method was established to produce isotope-labeled very-long-chain substrates, which can be used in future biochemical assays. The hypothetic substrate of CER1, C30 aldehyde, was also synthesized and used in preliminary in vivo assays of heterologously expressed CER1. No meaningful results were obtained from the assays, however, the developed protocols for substrate feeding and lipid extraction will be useful in future studies.  The major results of the previous chapters will be summarized in the following sections. Further research directions and possible experiments based on my results will also be discussed. 86  7.1 Synthesis of 1,2- and 1,3-bifunctional standard compounds Throughout this study, various long-chain and very-long-chain standard compounds with 1,2- and 1,3-bifunctionalities were synthesized to meet different identification needs. The 1,2-bifunctional standards synthesized include C8 and C14 α-ketols, C14 and C22 1,2-diol monoacetates (Chapter 3) and enantiomerically enriched C22 1,2-diols (Chapter 4). In Chapter 5, C20 and C30 β-keto acid methyl esters were synthesized, along with their derivatives, such as β-hydroxy acid methyl esters and alkan-2-ones. Only C20 β-keto NAC thioester was synthesized from the corresponding acyl-substituted Meldrum’s acid as a test of the reaction conditions. The obtained yields were acceptable (>70%) except for the asymmetric Sharpless dihydroxylations (12% and 40%) and the acid hydrolysis of β-functionalized acid methyl esters (51% and 58%). However, further optimizations of these reactions were not carried out, due to the purpose of this study.  Among the synthesized compounds, only 1,2-diol monoacetates, enantiomerically enriched 1,2-diols and β-hydroxy acid methyl esters were used directly in this thesis to identify either the structures or the stereoconfigurations of wax compounds. However, the other synthesized bifunctional compounds are still useful, as they can be used in the future as standards to identify unknown structures, or serve as substrates for biochemical assays. Furthermore, the established synthetic methods covered a large range of substrate chain lengths, and they can therefore be used in the future to produce standard compounds with different chain lengths when needed.  1,2- and 1,3-bifunctional compounds of structures different from those presented in this thesis may also be required in future research. Their synthesis can be built upon the currently available compounds. For example, the synthesis of enantiomerically enriched α-hydroxyacids or α-hydroxyaldehydes could be achieved by selectively oxidizing the primary alcohol groups of the corresponding 1,2-diols.129  87  7.2 Identification of novel 1,2- and 1,3-bifunctional wax compounds In Chapter 3, the two unknown compound series previously discovered from C. bipinnatus petal wax were identified as 1,2-alkanediol 1- and 2-acetates, with even-numbered chain lengths ranging from C20 to C24. Average ratios of 3:5 were found between 1,2-alkanediol 1- and 2-acetates , as opposed to the thermodynamic equilibrium ratio 7:3 of these two compounds. In Chapter 4, the unknown compound series found in A. arborescens leaf wax was identified as β-hydroxy acid methyl esters. The chain lengths of the β-hydroxy acid components range from C26 to C30, also dominated by even-numbered homologues.  To the best of my knowledge, only one example of wax compounds with 1,2-substitution pattern has been reported so far.130  Hence, the discoveries of 1,2-alkanediol 1- and 2-acetates and 1,2-alkanediols from C. bipinnatus petal wax provided another example of 1,2-bifunctional wax compounds. In addition, the novel structures of 1,2-alkanediol 1- and 2-acetates and β-hydroxy acid methyl esters have expanded current knowledge of possible wax compound structures.  Besides the structures mentioned above, it is very likely that other 1,2- and 1,3-bifunctional structures are also present in plant waxes, such as α-ketols and β-hydroxy acids. Identification of these structures can further enrich the structure pool of wax compounds. In fact, a search of novel β-functionalized wax compounds in Arabidopsis is currently underway. The hypothetic precursors of currently identified compounds, like α-hydroxy acid (see Chapter 4) and β-keto acid methyl ester (see Chapter 5) may also be present in plants, but in smaller amounts. Identification of these compounds could provide further evidence for possible biosynthetic pathways.  7.3 Determination of the stereoconfigurations of cosmos 1,2-diols Two independent NMR-based approaches, involving Mosher’s acid and the boronic acid, were established to study the stereoconfigurations of the secondary alcohol groups in 88  cosmos 1,2-diols. By comparing the NMR spectrum of 1,2-diols with those of enantiomerically enriched 1,2-diol standards, the stereocenters were found to have predominately the R-configuration (Chapter 4). An ee value of 62% of the cosmos 1,2-diols was determined using the boronic acid-based method.  The stereoconfigurations of wax compounds may carry important information of possible biosynthetic pathways, and therefore should be included in future structure identifications. The stereoconfigurations of other 1,2-bifunctional compounds, like 1,2-alkanediol 1- and 2-acetates, can be studied by first converting to 1,2-diols and then analyzing using the NMR-based methods established here. If the monoacetates have the same R-configurations as the 1,2-diols, this information can be used to further support their linkage in biosynthetic pathways. However, if the opposite results are obtained, it can be concluded that both compound groups originate from different pathways.  The Mosher’s acid method and the boronic acid-based NMR approaches can also be applied to 1,3-diols.84b, 131 However, enantiomerically enriched 1,3-diol standards need to be prepared first. To study the stereoconfigurations of other 1,3-bifunctional compounds, such as β-hydroxy acid methyl esters, they also need to be first reduced to 1,3-diols before analysis.  7.4 Synthesis of isotope-labeled substrates Both C20 and C30 β-deuterium substituted compounds were synthesized from the corresponding β-keto acid methyl esters via β-tosylate intermediates. The overall yields were 30% and 34%, respectively (Chapter 6). The C20 substrate was synthesized as mono-labeled compound, whereas the C30 substrate was double-labeled.  The percentage of the double-deuterium labeled compound in the obtained C30 substrate was determined to be 83%, and the remaining 17% was mono-labeled compound.  89  Two sets of conditions were developed for C20 and C30 substrates. Hence, compounds with chain lengths other C20 and C30 can also be synthesized using either of the two conditions. 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The Journal of Organic Chemistry 1991, 56 (2), 741-750.       106  Appendices Appendix A: Synthetic protocols of the compounds in Chapter 3  Octane-1,2-diol stannylene acetal 3-1: Dibutyltin oxide (2.02 g, 8.10 mmol) was added to a solution of 1, 2-diol octanediol (1.18 g, 8.10 mmol) in methanol (25 ml). The suspension was refluxed until it became clear. After an additional hour of refluxing, the solvent was evaporated and the residue was washed with hot acetone to give a white solid (3.0 g, 99% yield). 3-1 was used directly without further purification.   1-Hydroxy-octan-2-one 3-3: To a stirring solution of 3-1 (540.9 mg, 1.43 mmol) in 10 ml of CHCl3 at rt, was slowly added Br2 (275 mg / 88μl, in 5 ml CHCl3). The reaction was continued for 1 h after the addition of the Br2. At this point, most of the starting material had been consumed (monitored by TLC). The solvent was removed under vacuum, and the residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 4:1) to yield 3-3 as a white solid (154 mg, 75% yield).  1H-NMR (300 MHz, CDCl3): δ 4.23 (2H, s, CH2OH), 3.10 (1H, br s, -OH), 2.40 (2H, t, J=7.5 Hz, CH2CO), 1.63 (2H, m, CH2CH2CO), 1.10-1.40 (6H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 ppm (f1) 1.02.03.04.05.06.07.0 0 5000 10000 15000 107  Hz, -CH3).   Tetradecane-1,2-diol stannylene acetal 3-2: Dibutyltin oxide (1.62 g) was added to a solution of 1, 2-diol tetradecanediol (1.5 g) in methanol (25 ml), the suspension was refluxed until it became clear. After an additional hour of refluxing, the solvent was evaporated off, and the residue was washed with hot acetone to give a white solid (2.94 g, 98% yield).   1-Hydroxy-tetradecan-2-one 3-4: To a stirring solution of 3-2 (563.5 mg) in 10 ml of CHCl3 at rt, was slowly added Br2 (234.24 mg / 75μl, in 5 ml CHCl3). The reaction was continued for 1 h after the addition of Br2. Then the solvent was removed under vacuum, and the residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 6:1) to yield 3-4 as a white solid (224 mg, 80% yield).  1H-NMR (300 MHz, CDCl3): δ 4.24 (2H, s, CH2OH), 3.10 (1H, br s, -OH), 2.40 (2H, t, J=7.5 Hz, CH2CO), 1.63 (2H, m, CH2CH2CO), 1.10-1.40 (18H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).  ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 108   Tetradecane-1,2-diol monoacetate mixture (3-5 and 3-6): 2.4 mg of C14 diol was placed in an insert in GC vial and 2 microliters of acetic anhydride (2 equiv.) and 10 microliters of pyridine were injected into the vial insert. After staying at 70°C for 2 h, excess reagents were removed under nitrogen gas flow, followed by BSTFA/pyridine derivatization. Sample was diluted and analyzed by GC-MS.   Docosane-1,2-diol bis-acetate: 0.1 mg of C22 diol was placed in an insert in GC vial, and reacted with 10 microliters of acetic anhydride and 10 microliters of pyridine at 70°C for 2 h. Then dry the liquid under nitrogen gas flow, followed by BSTFA / pyridine derivatization. Sample was diluted and analyzed by GC-MS.   Docosane-1,2-diol monoacetate mixture (3-7 and 3-8): 0.5 mg of C22 diol was placed in an insert in GC vial, and reacted with 2 microliters of acetic anhydride (15 equiv.) and 10 microliters of pyridine for 0.5 h at rt. Then dry the liquid under nitrogen gas flow, followed by BSTFA / pyridine derivatization. Sample was diluted and analyzed by GC-MS.   1-Hydroxy-tetradecan-2-one 1-acetate 3-9: To a solution of 3-4 (30 mg) in 2 ml of distilled pyridine, acetic anhydride (67 mg) was slowly added. The mixture was stirred for 2 h at 50ºC, then quenched with water and extracted with ethyl acetate. The organic layer was 109  dried over anhydrous Na2SO4, and the solvent was removed under vacuum. Crude product was purified by TLC (silica, 2 mm layer, hexane: ethyl acetate 8:1) to afford 20 mg of 3-9 (yield = 58%).  1H-NMR (300 MHz, CDCl3): δ 4.65 (2H, s, COCH2OAc), 2.40 (2H, t, J=7.5 Hz, CH2CO), 2.17 (3H, s, OCOCH3), 1.60 (2H, m, CH2CH2CO), 1.10-1.40 (18H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   1,2-Tetradecanediol 1-acetate 3-5: 20 mg of 3-9 was dissolved in 2 ml of freshly distilled CHCl3 and 2 ml of distilled THF, and NaBH4 (67 mg) was added. The mixture was stirred at rt for 2 h, then quenched with diluted HCl. Extract the mixture twice with ethyl acetate. The combined organic phases were dried over anhydrous Na2SO4, and the solvent was removed under vacuum. Crude product was purified by TLC (silica, 2 mm layer, hexane: ethyl acetate 5:1) to give the desired product 3-5. ppm (f1) 1.02.03.04.05.06.07.08.0 0 1000 2000 3000 4000 110   1H-NMR (300 MHz, CDCl3): δ 4.15 (1H, dd, J=11.4, 3.0 Hz, CH(OH)CHHOAc), 3.95 (1H, dd, J=11.4, 7.4 Hz, CH(OH)CHHOAc), 3.84 (1H, m, CH(OH)CHHOAc), 2.10 (3H, s, OCOCH3), 1.47 (2H, m, CH2CH(OH)), 1.10-1.40 (18H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).  Detailed inspection of the TLC revealed a second product (3-6) characterized by clearly distinct signals in NMR.  1H-NMR (300 MHz, CDCl3): δ 4.91 (1H, m, CH(OAc)CHHOH), 3.72 (1H, N/A, CH(OAc)CHHOH), 3.64 (1H, N/A, CH(OAc)CHHOH), 2.09 (3H, s, CH(OCOCH3)), 1.45-1.65 (2H, m, CH2CH(OAc)), 1.10-1.40 (18H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 111  Appendix B: Synthetic protocols of the compounds in Chapter 4  (S)-Docosane-1,2-diol 4-1: Dissolve AD-mix-alpha (757.5 mg) in a t-BuOH/H2O 1:1 mixture (10 ml) and stirred at rt for 30 min. The mixture was cooled to 0°C and then 1-docosene (200 mg, 0.648 mmol) was added. Stir at rt overnight. There was still some starting material left according to TLC. Add 806 mg of Na2SO3 to the mixture. Stir at 0°C for 1h and extract with ethyl acetate. The combined organic phases were dried over anhydrous Na2SO4, and the solvent removed under vacuum. Crude product was subject to column chromatography (silica, gradient elution from pure hexane to pure ethyl acetate) to yield the product as a white solid (27 mg, 12% yield). 168 mg of unreacted 1-docosene was recovered.  1H-NMR (300 MHz, CDCl3): δ 3.73 (2H, m, CH(OH)CHHOH), 3.47 (1H, m, CH(OH)CHHOH), 1.10-1.90 (38H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   (R)-Docosane-1,2-diol 4-2: Dissolve AD-mix-alpha (781 mg) in a t-BuOH/H2O 1:1 mixture (10 ml) and stirred at room temperature for 30 min. The mixture was cooled to 0°C and then 1-docosene (150 mg, 0.486 mmol) was added. Stir at rt overnight. There was remaining starting material according to TLC. Add 806 mg of Na2SO3 to the mixture. Stir at 0°C for 1h and extract with ethyl acetate. The combined organic phases were dried over anhydrous Na2SO4, and the solvent removed under vacuum. Crude product was subject to ppm (f1) 1.02.03.04.05.06.07.0 0 1000 2000 3000 4000 5000 112  column chromatography (silica, gradient elution from pure hexane to pure ethyl acetate) to yield the product as a white solid (65.8 mg, 40% yield). 26 mg of unreacted 1-docosene was recovered.  1H-NMR (300 MHz, CDCl3): δ 3.73 (2H, m, CH(OH)CHHOH), 3.47 (1H, m, CH(OH)CHHOH), 1.10-1.90 (38H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).  Protocol of Mosher’s acid derivatization76 To a solution of R-1,2-docosanediol (2 mg, 0.0058 mmol) in chloroform, were added R-MTPA-Cl (4.4 mg, 0.017 mmol, 3 equiv.) and pyridine (1.64 μl, 0.020 mmol, 3.5 equiv.). The mixture was allowed to stir at rt overnight. The mixture was concentrated under vacuum and fractioned by TLC plate (silica, 0.5 mm, 20*20 cm, hexane: ethyl acetate 8:1). The major product (Rf = 0.26) was removed from TLC plate and extracted with fresh chloroform. The resulting product was subject to further analysis.  Derivatization protocol using boronic acid-based method85 To a solution of 1,2-tetradecanediol (9.22 mg, 0.04 mmol) in 2 ml of CDCl3, were added 2-formyl phenyl boronic acid 2-6 (6.00 mg, 0.04 mmol) and S-alpha-methyl benzylamine 2-7 (5.332mg, 5.672 μl, 0.044 mmol). The mixture was allowed to stir at rt for 15 min, and then an aliquot of the mixture was subject to NMR analysis.  ppm (f1) 1.02.03.04.05.06.07.08.0 0 1000 2000 3000 4000 5000 6000 7000 113  Appendix C: Synthetic protocols of the compounds in Chapter 5  Octadecanoyl meldrum’s acid 5-3: Dissolve stearic acid 5-1 (200 mg, 0.70 mmol), DCC (159 mg, 0.77 mmol), DMAP (90.2 mg, 0.74 mmol) in 5 ml distilled DCM. After 5 min, Meldrum’s acid 5-2 (101 mg, 0.703 mmol) was added to the solution. The mixture was allowed to stir overnight at rt and filtered through a short Celite column, eluding with DCM. Remove DCM under vacuum. The residue was taken up in ethyl acetate, washed with diluted HCl and dry over anhydrous Na2SO4. Remove solvent under vacuum afforded the product as a white solid (223 mg, yield = 77%).  1H-NMR (300 MHz, CDCl3): δ 3.06 (2H, t, J=7.5 Hz, CH2C(OH)), 1.73 (6H, s, C(CH3)2), 1.10-1.40 (30H, br m, aliphatic CH), 0.88 (3H, t, J=6.0 Hz, -CH3).   Methyl 3-oxoicosanoate 5-5: Dissolve 5-3 (223 mg, 0.54 mmol) in 7 ml of methanol and reflux for three hours. Remove solvent under vacuum. Residue was recrystallized from a mixture of hexane: methanol (2:1) to afford the pure product as a white solid (131.4 mg, yield = 71%). ppm (f1) 0.01.02.03.04.05.06.07.08.0 0 500 114   1H-NMR (300 MHz, CDCl3): δ 3.74 (3H, s, COOCH3), 3.44 (2H, s, COCH2COOCH3), 2.52 (2H, t, J=7.2 Hz, CH2CO), 1.10-1.70 (30H, br m, aliphatic CH), 0.88 (3H, t, J=6.4 Hz, -CH3).   S-(2-acetamidoethyl) 3-oxoicosanethioate 5-4: Dissolve N-acetyl cysteamine (51.2 mg, 0.426 mmol) in 2 ml of freshly distilled benzene, and add it dropwise to a stirring solution of 5-3 (262.4 mg, 0.64 mmol) in 3 ml of benzene. The mixture was refluxed overnight and then cooled to rt. Collect the precipitate by vacuum filtration and the filter cake was washed with benzene to afford the product as a white solid (143 mg, yield = 79%).  1H-NMR (300 MHz, CDCl3): δ 5.87 (1H, brs, NH), 3.69 (2H, s, COCH2CO), 3.46 (2H, q, J=6.1 Hz CH2N), 3.09 (2H, t, J=6.0 Hz SCH2), 2.52 (2H, t, J=7.2 Hz CH2CO), 1.98 (3H, s, ppm (f1) 1.02.03.04.05.06.07.08.0 0 1000 2000 3000 4000 5000 6000 7000 ppm (f1) 1.02.03.04.05.06.07.0 0 1000 2000 3000 4000 5000 115  NHCOCH3), 1.58 (2H, m, CH2CH2CO),1.18-1.38 (28H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   3-Oxoicosanoic acid 5-6: At rt, add 2 ml of acetic acid to 5-5 (56.9 mg, 0.167 mmol) and slightly warm it to dissolve the solid. Then carefully add concentrated HCl to the solution until precipitate started to persist. Add a few more drops of acetic acid to carefully adjust the solution to near transparent. Let the mixture stand for 3 day at rt. Collect the white solid by vacuum filtration. The crude product was taken up in chloroform, washed with NaHCO3 and dried over anhydrous Na2SO4. Remove the solvent under vacuum afforded the product as a white solid (28 mg, yield = 51%).  1H-NMR (300 MHz, CDCl3): δ 3.54 (2H, s, COCH2COOH), 2.56 (2H, t, J=7.5 Hz, CH2CO), 1.62 (2H, m, CH2CH2CO), 1.15-1.38 (28H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Nonadecan-2-one 5-7: In a round bottle, 5-5 (16 mg, 0.047 mmol) was dissolved in 2 ml of acetone and refluxed for 2 h. The solvent was removed under vacuum, and the residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 116  acetate 20:1) to yield 5-7 as a white solid (13 mg, yield = 98% ).  1H-NMR (300 MHz, CDCl3): δ 2.45 (2H, t, J=7.5 Hz, CH2COCH3), 2.16 (3H, s, CH2COCH3), 1.61 (2H, m, CH2CH2CO), 1.10-1.40 (28H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-hydroxyicosanoate 5-8: To a solution of 5-5 (100 mg, 0.294 mmol) in 2 ml of freshly distilled CHCl3 and 2 ml of distilled THF, was added NaBH4 (88 mg, 6 equiv.). The mixture was stirred at rt for 1.5 h and then quenched with diluted HCl. Extract the mixture with Chloroform. The combined organic layers were dried over Na2SO 4, filtered and concentrated under reduced pressure. The residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 10:1) to yield 5-8 as a white solid (76.3 mg, yield = 76% ). ppm (f1) 1.02.03.04.05.06.07.0 0 1000 2000 3000 4000 117   1H-NMR (300 MHz, CDCl3): δ 4.01 (1H, m, CH(OH)), 3.71 (3H, s, COOCH3), 2.52 (1H, dd, J=16.5, 3.3 Hz, CHHCOOMe), 2.40 (1H, dd, J=16.5, 8.7 Hz, CHHCOOMe), 1.10-1.70 (32H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   3-Hydroxyeicosanoic acid 5-9: At rt, 2 ml of acetic acid was added to 5-8 (20 mg, 0.058 mmol) placed in a glass vial. Slightly warm the vial to dissolve the solid. Then add concentrated HCl to the solution until precipitate started to persist. Add a few more drops of acetic acid to make the solution slightly opaque. Let the mixture stand for 3 day at room temperature and then collect the white solid by vacuum filtration. The crude product was taken up in chloroform, washed with NaHCO3 and dried over anhydrous Na2SO4. Remove solvent under vacuum afforded the product as a white solid (11 mg, yield = 58%).  ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 3000 3500 118   1H-NMR (300 MHz, CDCl3): δ 4.03 (1H, m, CH(OH)), 2.58 (1H, dd, J=16.5, 3.3 Hz, CHHCOOMe), 2.47 (1H, dd, J=16.5, 8.7 Hz, CHHCOOMe), 1.10-1.70 (32H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Octacosanoyl-Meldrum’s acid 5-11: Dissolve 5-10 (200 mg, 0.471 mmol), EDC (451 mg, 2.35 mmol) and DMAP (230 mg, 1.88 mmol) in 20 ml freshly distilled DCM. After stirring for 30 min at rt, Meldrum’s acid 5-2 (271 mg, 1.88 mmol) was added to the mixture and the mixture was allowed to stir overnight. The reaction mixture was filtered through a filtering paper under vacuum to get rid of precipitated urea. The organic phase was washed sequentially with diluted HCl and NaHCO3, and dried over anhydrous Na2SO4. The solvent was removed under vacuum, and the residue was taken up in ethyl chloroform and purified by column chromatography (silica, hexane: ethyl acetate 5:1) to give the product as a white solid (235 mg, yield = 91%). ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 119   1H-NMR (300 MHz, CDCl3): δ 3.06 (2H, t, J=7.2 Hz, CH2CO), 1.73 (6H, brs, C(CH3)2 ), 1.10-1.70 (50H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-oxotriacontanoate 5-12: Dissolve 5-11 (235 mg, 0.427 mmol) in 25 ml of methanol and reflux for eight hours. Then the solution was cooled to rt. Precipitate was collected by vacuum filtration to afford the product as a white solid (202 mg, yield = 98%).  1H-NMR (300 MHz, CDCl3): δ 3.74 (3H, s, COOCH3), 3.44 (2H, s, COCH2COOCH3), 2.52 (2H, t, J=7.2 Hz, CH2CO), 1.10-1.70 (50H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).  ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 ppm (f1) 1.02.03.04.05.06.07.0 0 500 1000 120   Nonacosan-2-one 5-13: To a solution of 5-12 (50 mg, 0.104 mmol) in 3 ml of DMF and 500 μl of water was added KOH (132 mg). The mixture was refluxed at 120°C for 4 h, and then quenched with diluted HCl. Extract the mixture with Chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 20:1) to afford the product as a white solid (31.5 mg, yield =73%).  1H-NMR (300 MHz, CDCl3): δ 2.41 (2H, t, J=7.5 Hz, CH2COCH3), 2.13 (3H, s, CH2COCH3), 1.10-1.90 (50H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Nonacosan-2-ol 5-14: To a solution of 5-13 (15 mg) in mixed CHCl3 and THF (2 ml, 1:1) was added NaBH4 (11 mg). The mixture was quenched with diluted HCl after stirring overnight at rt. Extract the product with chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was taken up in chloroform and purified by TLC plate (silica, 1mm, 20*20 cm, hexane: ethyl acetate 11:1) to afford the product as a white solid. (31.5 mg, yield =73%) ppm (f1) 1.02.03.04.05.06.07.08.0 0 1000 2000 3000 4000 121   1H-NMR (300 MHz, CDCl3): δ 3.80 (1H, m, CH(OH)), 1.20-1.50 (52H, br m, aliphatic CH), 1.18 (3H, d, J=6.3 Hz, CH(OH)CH3), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-hydroxytriacontanoate 5-15: To a solution of 5-12 (48 mg, 0.1 mmol) in 4 ml of distilled CHCl3 and 4 ml of distilled THF was added NaBH4 (48 mg, 0.1 mmol). The mixture was allowed to stir at rt overnight before quenched with diluted HCl. Extract the mixture with Chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was purified by recrystallization from MeOH to yield the product as a white solid (34.5 mg, yield = 72 %).  1H-NMR (300 MHz, CDCl3): δ 4.01 (1H, m, CH(OH)), 3.71 (3H, s, COOCH3), 2.52 (1H, dd, J=16.2, 3.0 Hz, CHHCOOMe), 2.40 (1H, dd, J=16.2, 8.7 Hz, CHHCOOMe), 1.10-1.70 (52H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).  ppm (f1) 1.02.03.04.05.06.07.0 0 500 1000 1500 2000 2500 3000 ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 3000 3500 122  Appendix D: Synthetic protocols of the compounds in Chapter 6  Methyl 3-iodoeicosanoate 6-1: A 25 ml round bottle was charged with PPh3 (100.7 mg, 0.384 mmol), imidazole (24.1 mg, 0.384 mmol), iodine (73 mg, 0.288 mmol) and 5-8 (32.5 mg, 0.096 mmol). 4.5 ml distilled toluene was added to the round bottle and the mixture was heated to reflux. TLC showed complete consumption of the starting material after 2 h. 5 ml of diluted Na2SO3 was add to quench the reaction and the mixture was extracted with ether. The combined organic layers were dried over Na2SO4, filtered and solvent was removed under reduced pressure. Crude product was purified with TLC (20*20 cm, 1 mm layer, hexane: ethyl acetate 15:1) and then with column chromatography (silica, hexane: ethyl acetate 15:1). A side product with similar Rf value could not be separated. The NMR of the crude product is as shown.  1H-NMR (300 MHz, CDCl3): δ 4.35 (1H, m, CH(I)), 3.72 (3H, s, COOCH3), 3.0 (2H, d, J=7.2 Hz, CHHCOOMe), 1.10-1.70 (32H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   (E)-methyl eicos-2-enoate 6-2: Dissolve the crude product from last step in 2 ml distilled THF, then LiAlH4 (large excess) was added. The mixture turned dark yellow upon addition ppm (f1) 1.02.03.04.05.06.07.08.0 0 100 200 300 400 500 600 700 123  of LiAlH4 and then gradually became light yellow. After 1 hour of reflux, TLC showed complete conversion of the starting material. The reaction was quenched with crushed ice and diluted HCl after additional 0.5 h of reflux. The reaction mixture was extracted with ether. The combined organic layers were dried over Na2SO4, filtered and the solvent was removed under reduced pressure. The residue was purified with column chromatography (silica, hexane: ethyl acetate 10:1) to afford the product. According to NMR and GC-MS results, the product was characterized as methyl eicos-2-enoate (elimination product). In the iodination step, elimination also occurred, according to NMR.  1H-NMR (300 MHz, CDCl3): δ 6.98 (1H, m, CH=CHCOOMe), 5.82 (1H, d, J=15.9 Hz, CH=CHCOOMe), 3.72 (3H, s, COOCH3), 2.2 (2H, m, CH2CH=CH), 1.10-1.70 (32H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-(tosyloxy)eicosanoate 6-3: Dissolve 5-8 (136.9 mg, 0.404 mmol) and p-toluenesulfonyl chloride (154.2 mg, 0.809 mmol) in 7 ml of distilled DCM. Then DMAP (99 mg, 0.809 mmol) and triethylamine (113 ul, 0.809 mmol) were added to the stirring mixture at rt. Let the reaction stir overnight and then quench the reaction with diluted HCl. The mixture was extracted with Chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was purified by TLC plate (silica, 2mm, 20*20 cm, hexane: ethyl acetate 7:1) to yield the product as a white ppm (f1) 1.02.03.04.05.06.07.0 0 50 100 150 200 124  solid (95 mg, yield = 48 %).  1H-NMR (300 MHz, CDCl3): δ 7.80 (2H, d, J=8.2 Hz, ArH), 7.33 (2H, d, J=8.2 Hz, ArH), 4.86 (1H, pentet, CH(OTs)), 3.61 (3H, s, COOCH3), 2.76 (1H, dd, J=15.6, 6 Hz, CHHCOOMe), 2.57 (1H, dd, J=15.9, 6.9 Hz, CHHCOOMe), 2.44 (3H, s, CH3Ar), 1.61 (2H, m, CH2CH(OTs)), 1.10-1.70 (30H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-deuterium-eicosanoate 6-4: 1 ml of DMSO was added to a mixture of 6-3 (5.4 mg, 0.011 mmol) and NaBD4 (large excess) in a 5 ml round bottle. The reaction was stirred at 70°C for 3.5 h and then quenched with diluted HCl and water. The reaction mixture was extracted with hexane. The combined organic layers were dried over Na2SO4, filtered and the solvent was removed under reduced pressure to afford the product as a white solid (3 mg, yield = 83%). ppm (f1) 1.02.03.04.05.06.07.08.09.0 0 500 1000 1500 2000 125   1H-NMR (300 MHz, CDCl3): δ 3.66 (3H, s, COOCH3), 2.29 (2H, d, J=7.8 Hz, CH2COOMe), 1.10-1.70 (35H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-deuterium-3-hydroxytriacontanoate 6-5: Dissolve 5-12 (140 mg, 0.291 mmol) in 4 ml of distilled CHCl3 and 4 ml of distilled THF, then NaBD4 (60 mg, 1.45 mmol) was added to the solution. The mixture was stirred at rt overnight and then quenched with diluted HCl. The mixture was extracted with Chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was purified by recrystallization from MeOH to yield the product as a white solid (130 mg, yield = 92 %).  1H-NMR (300 MHz, CDCl3): δ 3.71 (3H, s, COOCH3), 2.52 (1H, d, J=16.5 Hz, CHHCOOMe), 2.40 (1H, d, J=16.5 Hz, CHHCOOMe), 1.10-1.70 (52H, br m, aliphatic CH), ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 3000 ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 3000 3500 126  0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3-deuterium-3-(tosyloxy)triacontanoate 6-6: Dissolve 6-5 (103 mg, 0.213 mmol) and p-toluenesulfonyl chloride (256 mg, 1.34 mmol) in 3 ml of distilled DCM. Then DMAP (164 mg, 1.34 mmol) and triethylamine (188 μl, 1.34 mmol) were added to the stirring mixture at rt. Let the reaction stir for 3.5 h and then quench the reaction with diluted HCl. The mixture was extracted with Chloroform. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was purified by TLC plate (silica, 2mm, 20*20 cm, hexane: ethyl acetate 7:1) to yield the product as a white solid (86.8 mg, yield = 64 %).  1H-NMR (300 MHz, CDCl3): δ 7.80 (2H, d, J=8.1 Hz, ArH), 7.33 (2H, d, J=7.8 Hz, ArH), 3.61 (3H, s, COOCH3), 2.77 (1H, d, J=15.6 Hz, CHHCOOMe), 2.57 (1H, d, J=15.6 Hz, CHHCOOMe), 2.44 (3H, s, CH3Ar), 1.62 (2H, m, CH2CH(OTs)), 1.10-1.70 (50H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Methyl 3,3-dideuteriumtriacontanoate 6-7: To a mixture of 6-5 (84.7 mg, 0.133 mmol) and NaBD4 (43.6 mg, 1.06 mmol) in a 25 ml round bottle was added 5 ml of DMSO. The ppm (f1) 1.02.03.04.05.06.07.08.0 0 500 1000 1500 2000 2500 127  reaction was allowed to stir at 100°C for 3.5 h. Quench the reaction with diluted HCl and further dilute it with water. The mixture was then extracted with hexane. The combined organic layers were dried over Na2SO4, filtered and concentrated under reduced pressure. The residue was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 20:1) to afford the product as a white solid (36.0 mg, yield =58%).  1H-NMR (300 MHz, CDCl3): δ 3.66 (3H, s, COOCH3), 2.28 (2H, s, CD2CH2COOMe), 1.10-1.90 (52H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Stearaldehyde 6-9: To a stirring suspension of pyridinium chlorochromate (PCC, 537 mg, 1.3 equiv.) in methlyene chloride (CH2Cl2, 20 ml) was added 1-octadecanol (503 mg, 1.86 mmol in 5 mL of CH2Cl2). The reaction was allowed to stir at rt. 30 ml of ethyl ether was added when TLC showed no presence of starting material (1.2 h). Decant the supernatant and the insoluble residual was washed three times with ether. The combined organic solution was passed through a short pad of Celite, and the filtrate was concentrated under vacuum to yield the crude product as a white solid. Crude product was taken up in hexane and purified by column chromatography (silica, hexane: ethyl acetate 30:1) to yield 1-octadecanal as a white solid (334.2 mg, yield = 67%). ppm (f1) 1.02.03.04.05.06.07.08.0 0 100 200 300 400 500 128   1H-NMR (300 MHz, CDCl3): δ 9.76 (1H, t, J=1.8 Hz, CHO), 2.42 (2H, td, J=7.3, 1.8 Hz, CH2CHO), 1.64 (2H, m, CH2CH2CHO), 1.10-1.40 (48H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3).   Triacontanal 6-11: To a refluxing suspension of pyridinium chlorochromate (PCC, 20.6 mg, 0.0955) in methlyene chloride (CH2Cl2, 4 ml) was added 1-triacontanol (15.2 mg, 0.0344 mmol, in 2 ml CH2Cl2). After 2 h, TLC showed no presence of starting material. The mixture was directly subject to column chromatography (silica, hexane: ethyl acetate 20:1) to yield 1-triacontanal as a white solid (7.0 mg, yield = 46%).  1H-NMR (300 MHz, CDCl3): δ 9.76 (1H, t, J=1.8 Hz, CHO), 2.42 (2H, td, J=7.3, 1.8 Hz, CH2CHO), 1.64 (2H, m, CH2CH2CHO), 1.10-1.40 (48H, br m, aliphatic CH), 0.88 (3H, t, J=6.9 Hz, -CH3) ppm (f1) 0.05.010.0 0 500 1000 1500 2000 2500 ppm (f1) 5.010.0 0 100 200 300 400 500 600 700

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