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Investigating the biosynthesis of cuticular alkanes in Arabidopsis thaliana : Characterization of SCD2… Skvortsova, Mariya 2013

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INVESTIGATING THE BIOSYNTHESIS OF CUTICULAR ALKANES IN ARABIDOPSIS THALIANA: CHARACTERIZATION OF SUSCEPTIBLE TO CORONATINE DEFICIENT PSEUDOMONAS SYRINGAE 2  by  Mariya Skvortsova B.A., Rutgers University, 2010  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE  in  The Faculty of Graduate Studies  (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2013 © Mariya Skvortsova, 2013  Abstract	
   The plant cuticle is a hydrophobic layer that seals the surface of primary aerial organs of terrestrial plants and serves in protecting the tissues from abiotic and biotic stresses. Lipids are synthesized in the plastid and in the endoplasmic reticulum (ER) of epidermal cells for eventual export and deposition on the surface. Great progress has been made by genetic studies in the model plant Arabidopsis thaliana in elucidating fatty acid elongation, but knowledge of alkane biosynthesis is still scarce. The current work was focused on expanding our current understanding of alkane biosynthesis in Arabidopsis thaliana. A recent discovery of Susceptible to Coronatine-Deficient Pst DC3118-2 (SCD2) whose mutant has a leaf-specific increase in aldehydes and a decrease in alkanes suggests that SCD2 has a role in converting aldehydes to alkanes. In this thesis, further characterization of SCD2 revealed that alkanes are decreased in two mutant lines, the wax of mutants was restored by transgene complementation with the native gene, the transcript is abundant in leaves, and the promoter is active in the phloem of vasculature. Finally, the protein localized to the ER, consistent with its role in wax biosynthesis. This work provided evidence for yet another gene whose product is involved in formation of cuticular alkanes in Arabidopsis thaliana. Double mutants were generated to further study wax biosynthesis in both stems and leaves. The cer1cer3 mutant had greatly reduced total stem and leaf wax amounts compared to wild-type, as well as a substantial reduction of alkanes. It has an increase in C30 primary alcohol levels like the cer3 parent, indicating epistasis. This suggests that CER3 precedes CER1 in alkane formation. Furthermore, it is severely male-sterile with a reduction in epicuticular wax crystals. Wax biosynthesis is similar in stems and leaves of cer1cer3, cer1cer4 and cer3cer4. The cer1cer3 will be an important tool to test domain functionality of CER1 and CER3 and may shed more light on the mechanisms of alkane formation in Arabidopsis thaliana.  	
    ii	
    Preface	
   The work presented in Chapter 3 was in partial collaboration with Dr. Weiqing Zeng. He sent homozygous scd2-1 and scd2-2 seeds, along with the respective Col-7 and Ler ecotypes. He also provided the pSCD2:SCD2-GFP in scd2-1 complementation line. I have extracted the wax from all reported lines and analyzed the mass spectra. I extracted RNA from Col-0, synthesized cDNA and designed gene-specific primers for qRT-PCR analyses of SCD2 gene expression. I cloned the p35S:SCD2-GFP and pSCD2:GUS constructs, transformed them into Arabidopsis scd2-1 and Col-0, respectively and performed all the confocal microscopy (with assistance of Sandra Keerthisinghe) and histochemical staining myself.  In Chapter 4, I have performed the crossing myself, and genotyped and analyzed single and double mutant lines. All experiments were performed by me.  The constructs harboring truncations and fusions of CER1 and CER3 domains were cloned by me and will serve as material for future experiments. I also wrote the manuscript of this thesis. Thesis editing was performed with the help of my supervisor Dr. Jetter.  	
    iii	
    Table of Contents Abstract.......................................................................................................................................... ii Preface ........................................................................................................................................... iii Table of Contents ......................................................................................................................... iv List of Tables ............................................................................................................................... vii List of Figures............................................................................................................................. viii List of Abbreviations ................................................................................................................... xi Acknowledgements .................................................................................................................... xiii Dedication ................................................................................................................................... xiv Chapter 1 Introduction to plant cuticles..................................................................................... 1 1.1 Ecological and physiological roles of plant cuticles ......................................................... 1 1.2 Structure of plant cuticles .................................................................................................. 1 1.3 Composition of plant cuticles ............................................................................................. 2 1.4 Wax biosynthesis-fatty acid elongation............................................................................. 3 1.5 The biosynthesis of wax compounds ................................................................................. 6 1.5.1 Synthesis of primary alcohols and wax esters ............................................................... 6 1.5.2 Synthesis of alkanes ....................................................................................................... 7 1.5.3 Synthesis of secondary alcohols and ketones .............................................................. 12 1.6 Research questions and objectives .................................................................................. 12 Chapter 2 Materials and methods ............................................................................................. 19 2.1 Plant materials and growth conditions ........................................................................... 19 2.2 Wax analyses ..................................................................................................................... 19 2.2.1 Wax extraction and derivatization ............................................................................... 19 2.2.2 Chemical analyses using GC-FID and GC-MS ........................................................... 20 2.3 Genomic DNA extraction, RNA isolation, cDNA synthesis and reverse transcription ................................................................................................................................................... 20 2.3.1 Genomic DNA extraction for PCR genotyping ........................................................... 20 2.3.2 RNA and cDNA preparation, quantitative RT-PCR conditions and analysis ............. 21 2.4 Scanning electron microscopy (SEM) ............................................................................. 21 	
    iv	
    2.5 Cloning and subcellular localization of SCD2 in Arabidopsis thaliana and Nicotiana benthamiana ............................................................................................................................. 22 2.5.1 Transient expression of SCD2 in Nicotiana benthamiana ........................................... 24 2.5.2 Light microscopy and laser scanning confocal microscopy ........................................ 24 2.5.3 Promoter-GUS cloning and histochemical staining ..................................................... 25 Chapter 3 Molecular characterization of SCD2 ...................................................................... 27 3.1 Introduction ....................................................................................................................... 27 3.2 Results ................................................................................................................................ 29 3.2.1 Total stem wax ............................................................................................................. 30 3.2.2 Total leaf wax .............................................................................................................. 31 3.2.3 Organ-specific expression of SCD2 ............................................................................. 33 3.2.4 Tissue-specific expression of SCD2 ............................................................................ 34 3.2.5 Subcellular localization of SCD2................................................................................. 35 3.2.6 Transient expression of SCD2 in Nicotiana benthamiana .......................................... 35 3.2.7 Stable transformation of p35S:SCD2-GFP in Arabidopsis scd2-1 mutants ................ 36 3.3 Discussion .......................................................................................................................... 36 Chapter 4 Characterization of cer1cer3 and elucidation of epistatic relationships of CER genes in Arabidopsis wax biosynthesis ....................................................................................... 56 4.1 Introduction ....................................................................................................................... 56 4.2 Results ................................................................................................................................ 58 4.2.1 Generating the cer1cer3 double mutant ....................................................................... 58 4.2.2 The cer1cer3 double mutant exhibits a glossy stem phenotype .................................. 59 4.2.3 The cer1cer3 double mutant exhibits a severe additive sterility defect....................... 59 4.2.4 The cer1cer3 double mutant has a reduction of wax crystals ...................................... 59 4.2.5 Arabidopsis cer1, cer3 and cer1cer3 show altered cuticular wax composition .......... 60 4.2.6 Arabidopsis cer1cer4 and cer3cer4 show altered cuticular wax composition ............ 61 4.3 Discussion .......................................................................................................................... 63 4.3.1 The cer1cer3 double mutant is severely male-sterile .................................................. 64 4.3.2 Wax crystals are present on cer1cer3 .......................................................................... 66 4.3.3 CER3 precedes CER1 in both stems and leaves .......................................................... 66 4.3.4 The cer1cer3, cer1cer4 and cer3cer4 double mutants reveal complex gene interactions ............................................................................................................................................... 68 	
    v	
    4.3.5 Wax biosynthesis in stems and leaves ......................................................................... 69 Chapter 5 Summary and future directions .............................................................................. 83 5.1 Molecular characterization of SCD2 and accumulation of aldehydes in leaves of SCD2 mutants .......................................................................................................................... 84 5.2 Characterization of cer1cer3 and elucidation of epistatic relationships of CER genes in Arabiopsis wax biosynthesis ............................................................................................... 86 References .................................................................................................................................... 88  	
    	
    vi	
    List	
  of	
  Tables	
   Table 1 Cuticular wax composition of rosette leaves of Arabidopsis Col-7 (WT), scd2-1, Ler (WT), scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3)............. 48 Table 2 Cuticular wax composition of inflorescence stems of Arabidopsis Col-0 (WT), cer1, cer3, cer4, cer1cer3, cer1cer4 and cer3cer4. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3)............................................. 75 Table 3 Cuticular wax composition of rosette leaves of Arabidopsis Col-0 (WT), cer1, cer3, cer4, cer1cer3, cer1cer4 and cer3cer4. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3). ............................................................... 76  	
    vii	
    List	
  of	
  Figures	
   Figure 1 Scheme showing a cross-section of a plant cuticle and epidermal cells. Figure modified after Jetter et al (2000) and Jeffree (1996). ................................................................................... 15 	
   Figure 2 Wax Biosynthetic Pathway in Arabidopsis thaliana stems. CER, eceriferum; WSD, wax synthase/diacylglycerol acyltransferase; MAH, mid-chain alkane hydroxylase. Figure modified from Samuels et al (2008). ............................................................................................................ 16 	
   Figure 3 Trans-membrane prediction results of CER1 and CER3 presented by the TMHMM program. ........................................................................................................................................ 17 	
   Figure 4 Scheme of CER1 and CER3 proteins with desaturase and dehydrogenase (SDR) domains illustrated. Histidine-rich motifs are represented with orange boxes. The following scheme of domains was predicted using the hydropathy results of the TMHMM program......... 18 	
   Figure 5 Cuticular wax composition of leaves and stems of Arabidopsis Col-7 (WT) and scd2-1. Wax coverage is expressed as (µg/cm2). Mean values are given with SD (n=3). Each wax constituent is designated by carbon chain length and is labeled by chemical class along the xaxis. Taken from Zeng and He (2010). Abbreviations Alk=alkane, PA=primary alcohol, ALD=aldehyde, FFA=free fatty acid, KE=ketone........................................................................ 43 	
   Figure 6 Cutin monomer composition of leaves and stems of Arabidopsis Col-7 (WT) and scd21. Monomer amounts are expressed as (µg/cm2) of stem and leaf surface. Mean values are given with SD (n=3). Adapted from Zeng and He (2010). ..................................................................... 44 	
   Figure 7 Cuticular wax composition on inflorescence stems of Col-7 (WT), scd2-1, Ler, scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major compound classes are expressed as (% of total stem wax) of stem surface area. The data represent mean values ± SD of n=5. ..................................................................................................................................... 45 	
   Figure 8 Cuticular wax composition on inflorescence stems of Col-7 (WT), scd2-1, Ler, scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major components are expressed as (% of compound class) of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=5. ...................................................................................................................... 46 	
   Figure 9 Cuticular wax coverage on rosette leaves of Col-7 (WT), scd2-1, Ler (WT), scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Total wax coverage is expressed as µg/dm2 of leaf surface area. The data represent mean values ± SD of n=5.............................. 47 	
   Figure 10 Cuticular wax composition on rosette leaves of Col-7 (WT), scd2-1, Ler (WT), and scd2-2 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the xaxis. The data represent mean values ± SD of n=5. br=branched primary alcohols..................... 49 	
   Figure 11 Cuticular wax composition on rosette leaves of Col-7 (WT), scd2-1, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major components are expressed as 	
    viii	
    µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=5. br=branched primary alcohols. ..................................................................................................... 50 	
   Figure 12 Differential expression analysis of Arabidopsis SCD2 in various organs of four-weekold Arabidopsis plants. The gene expression level was determined by real-time RT-PCR analysis. Results are represented as relative transcript abundances. The data represent the means ± SD of three biological and two technical replicates (n=6). Total RNA was isolated from roots, rosette and cauline leaves, bottom- mid- and top section of stem, flowers and siliques. ............. 51 	
   Figure 13 Spatial expression pattern the SCD2 gene in transgenic Arabidopsis plants harboring the SCD2 promoter fused to the GUS gene. Promoter activity was visualized through histochemical GUS staining on two-week old T2 transgenic seedlings. A and B. Whole seedling imaged by dissecting light microscope, C and D. mature leaves, E. hypocotyl F. petiole and shoot apex, G. emerging true leaf, H. root tip and differentiation zone, I. secondary roots, J and K. DIC image of root, L. True leaf mid-vein. A-B bars= 5mm, J and L bars=20 µm, K bar=15 µm ,C-I bars=0.2 mm. ................................................................................................................................ 52 	
   Figure 14 Hydrophobicity plots of various wax biosynthetic proteins. A. SCD2 B. CER8 C. CER2 D. CER6. Data generated using amino acid sequences of various proteins in the TMHMM program. ........................................................................................................................................ 53 	
   Figure 15 Transient expression of p35S:SCD2-GFP and p35S:HDEL-GFP in N. benthamiana epidermal cells. A. p35S:SCD2-GFP, B. p35S:HDEL-GFP, and C. as merge. Bars=2µm. ........ 54 	
   Figure 16 Subcellular localization of SCD2 in root epidermal cells of stably transformed Arabidopsis scd2-1. Confocal microscopy images of p35S:SCD2-GFP. A. Z stack of root. B. root tip. C. root hairs and root epidermal cells. D, E, F, G. Planes through the root epidermal cell. H. root epidermal cell under 488 nm excitation (GFP). I. hexyl rhodamine stain under 561 nm excitation (RFP). J. merge. Bars=20µm A-G. .............................................................................. 55 	
   Figure 17 Strategy used to cross cer1 with cer3 to produce the cer1cer3 double mutant. A. The successful and B. unsuccessful crossing combinations. C. Genotyping of cer1, cer3, cer4 single mutants and cer1cer3, cer1cer4 and cer3cer4 double mutants. Gene specific primers were used to PCR-amplify the WT locus of CER1, CER3 and CER4. Primers are denoted by abbreviations F=forward primer, R=reverse primer, C1=CER1, C3=CER3, C4=CER4. The Lba1 primer was used in combination with the appropriate forward or reverse primers of the WT locus to check presence of the T-DNA in both single and double mutants. ......................................................... 72 	
   Figure 18 Phenotype characterization of the cer1cer3 mutant. A. Visual stem phenotype of Col0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant. B. Visual silique phenotype of Col-0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant. C. Quantitative analysis of sterility by seed count (per plant) from Col-0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant. ............................................................................ 73 	
   Figure 19 Scanning electron micrographs of wild-type, cer1-2, cer3-6, and cer1cer3 stems. Bars=2µm. ..................................................................................................................................... 74 	
   	
    ix	
    Figure 20 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer1, cer3 and cer1cer3 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. .................................................................. 77 	
   Figure 21 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer1, cer4 and cer1cer4 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. .................................................................. 78 	
   Figure 22 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer3, cer4 and cer3cer4 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. .................................................................. 79 	
   Figure 23 Cuticular wax composition on rosette leaves of Col-0 (WT), cer1, cer3 and cer1cer3 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the xaxis. The data represent mean values ± SD of n=3. ...................................................................... 80 	
   Figure 24 Cuticular wax composition on rosette leaves of Col-0 (WT), cer1, cer4 and cer1cer4 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the xaxis. The data represent mean values ± SD of n=3. ...................................................................... 81 	
   Figure 25 Cuticular wax composition on rosette leaves of Col-0 (WT), cer3, cer4 and cer3cer4 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the xaxis. The data represent mean values ± SD of n=3. ...................................................................... 82	
   	
   	
    	
    x	
    List	
  of	
  Abbreviations	
   ACP  acyl carrier protein  bp  base pair  BiFC  bimolecular fluorescence complementation  BSTFA  bis-N,O-(trimethylsilyl)trifluoroacetamide  CER  eceriferum  CoA  coenzyme A  ECR  enoyl-CoA reductase  ER  endoplasmic reticulum  FAE  fatty acid elongase  FAR  fatty acyl reductase  FID  flame ionization detection  GC  gas chromatography  GFP  green fluorescent protein  GUS  β-glucuronidase  HCD  β-hydroxyacyl-CoA dehydratase  KCR  β-ketoacyl-CoA reductase  KCS  β-ketoacyl-CoA synthase  LACS  long chain acyl-CoA synthetase  LTP  lipid transfer protein  MAH  mid-chain alkane hydroxylase  MS  mass spectrometry  p  promoter  PCR  polymerase chain reaction  	
    xi	
    q  quantitative  RFP  red fluorescent protein  RT  reverse transcriptase  SCD2  Susceptible to Coronatine-Deficient Pst DC3118-2  UV  ultraviolet  VLC  very-long-chain  VLCFA  very-long-chain fatty acid  WS/DGAT  wax synthase/diacylglycerol acyl transferase  X-Gluc  5-bromo-4-chloro-3-indolyl glucuronide  	
    xii	
    Acknowledgements	
   It is my pleasure to express my deepest appreciation and regards to my research supervisor, Dr. Reinhard Jetter, for his immeasurable support, instruction and supervision throughout my studies in his laboratory. This project would not have been possible without his valuable input and guidance. My further gratitude and warm thanks extend to the members of my supervisory committee Dr. Shawn Mansfield and Dr. Carl Douglas, whose invaluable advice and time helped steer my project in the successful direction and to completion.  I would like to thank my colleagues in the Jetter lab, including Dr. Christopher Buschhaus, Chen Peng, Ruonan Yao, Yan Cao, Luke Busta, Radu Racovita, Alvaro Luna, and Daniela Hegebarth for their extremely helpful technical advice and sharing constructive input. The friendly working atmosphere could not have been possible without them. In addition, my appreciation goes out to Dr. Mathias Schuetz for providing the p35S:HDEL-RFP marker and expertise with transient expression in Nicotiana benthamiana, as well as constructive advice. Also, thanks to Lin Shi for technical help and Sandra Keerthisinghe for her time and expertise with confocal microscopy. Finally, I thank Dr. Weiqing Zeng for providing seeds of SCD2 mutants and pSCD2:SCD2 in scd2-1 transgenic lines.  Finally, I would like to express my immense gratitude to my beloved parents whose undying support and encouragement have helped me immeasurably.  	
    xiii	
    Dedication	
    To my beloved mother and father  	
    xiv	
    Chapter	
  1	
  Introduction	
  to	
  plant	
  cuticles	
   1.1	
  Ecological	
  and	
  physiological	
  roles	
  of	
  plant	
  cuticles	
   The plant cuticle is a hydrophobic layer that seals the surface of primary aerial organs (Riederer and Schreiber, 1995). Its presence is essential for protecting plants against both abiotic and biotic stresses. The primary function of the cuticle is to prevent uncontrolled non-stomatal water loss (Stiles, 1994) and as such was instrumental for the adaptation and evolution of land plants (Raven and Edwards, 2004). The cuticle also promotes surface cleanliness (Kerstiens, 1996; Barthlott and Neinhuis, 1997; Riederer and Schreiber, 2001), deters against pathogen attack (Jenks et al., 2002) and herbivory (Eigenbrode and Espelie, 1995), and protects against ultraviolet (UV) radiation. Furthermore, the cuticle has a crucial role in plant development as it prevents the cell wall fusions between adjacent organs (Sieber et al., 2000; Javelle et al., 2011). 	
   In this introductory chapter, the structure and chemical composition of the plant cuticle is described in sections (1.2) and (1.3), respectively. Subsequently, the wax biosynthetic pathway will be discussed first by focusing on fatty acid elongation (section 1.4), followed by synthesis of primary alcohols and wax esters (1.5.1), alkanes (1.5.2) and secondary alcohols and ketones (1.5.3). The research questions and objectives are outlined in section 1.6. 	
    1.2	
  Structure	
  of	
  plant	
  cuticles	
   The cuticle is a multi-layered structure (Figure 1) (Jeffree, 1996). The outermost layer is a continuous and thin epicuticular wax film. Many species also have wax crystals that form on the film and project into the surrounding environment (Jetter et al., 2006). Beneath the epicuticular wax is the next layer consisting of intracuticular wax embedded into a cutin/cutan biopolymer matrix (Pollard et al., 2008). It is believed that cellulose fibrils likely extend into the cutin and 	
    1	
    intracuticular wax from the epidermal cell wall. However, it is not certain to what extent the fibrils project into the cuticle and their universal presence. A pectinaceous layer is thought to divide the cell wall from the cuticular layer in some species. However, the composition and species distribution of pectin are still uncertain.  1.3	
  Composition	
  of	
  plant	
  cuticles	
   The cuticular biopolymer matrix contains cutin and cutan, contributing 40-80% to the cuticle’s total weight (Heredia, 2003). Cutin consists of ω- and mid-chain hydroxy and epoxy C16 to C18 fatty acids linked by ester bonds (Heredia, 2003). The discovery of glycerol bridges was quite unexpected in cutin (Graça et al., 2002). In contrast, cutan contains cross-linking ether bonds that prevent it from depolymerizing upon ester hydrolysis. Consequently, a residue is left behind after cutin monomers have been depolymerized (Pollard et al., 2008; Beisson et al., 2012). Interestingly, Arabidopsis has an unusual cutin composition, enriched in dicarboxylic acids resembling the root-associated aliphatic polymer suberin (Bonaventure et al., 2004; Franke et al., 2005). The exact composition and structure of cutan must still be elucidated. The presence of cutin and cutan, as well as the ratio of the two is dependant on developmental stages and plant species.  On the other hand, cuticular waxes appear as a complex mixture, which can be readily extracted from the surface by organic solvents of low polarity (Jetter et al., 2006). Cuticular wax consists of aliphatics, alicyclics and aromatics (Jetter et al., 2006). The aliphatics predominate in cuticular waxes and consist of very-long-chain fatty acids (VLCFA) and their derivatives of chain lengths ranging from C20 to C34. The ubiquitous wax constituents are primary alcohols, wax esters, alkanes, and aldehydes (Jetter et al., 2006). The chain lengths of esters range from C38 to C70. Additionally, alicyclics and aromatics, mostly triterpenoids and phenolics, respectively are often 	
    2	
    found in minor amounts (Jetter et al., 2006). Triterpenoids consist of 30 carbons and are usually pentacyclic. The cyclic compounds along with very-long-chain (VLC) primary alcohols preferably accumulate in the intracuticular layer of the cuticle, whereas VLC fatty acids and alkanes do so in the epicuticular layer (Buschhaus and Jetter, 2011). Substantial differences of aliphatic chain length profiles were not observed between the epi- and intracuticular wax layers. Wax load and composition varies among plant species (Post-Beittenmiller, 1996). Furthermore, wax composition varies between different organs, tissues and developmental stages (Jenks et al., 1995; Rashotte et al., 1997; Jetter et al., 2001; Shepherd and Wynne Griffiths, 2006; Pollard et al., 2008; Li and Beisson, 2009; Van Maarseveen et al., 2009; Buschhaus and Jetter, 2011). A multiplicity of environmental factors, such as light, moisture and temperature can affect wax deposition during organ growth (Kolattukudy, 1996; Schreiber et al., 2001).  1.4	
  Wax	
  biosynthesis-­‐fatty	
  acid	
  elongation	
   VLCFAs are one of the constituents of waxes, and they serve as precursors for other wax constituents like primary alcohols, esters, alkanes, and secondary alcohols. VLCFAs are synthesized by the elongation of long-chain fatty acyl-CoAs.  The process starts with de novo synthesis of 16 and 18 carbon-long acyl chains by the soluble fatty acid synthase complexes (FAS) in the plastid stroma of epidermal cells. Prior to exiting the plastidial stroma, long-chain fatty acids need to be liberated from the acyl carrier protein (ACP) by an acyl-ACP thioesterase, and then exported from the plastid via an unknown mechanism (Bonaventure et al., 2003; Kunst et al., 2006). The C16 and C18 acids are then esterified to coenzyme A (CoA) to yield C16 and C18 acyl-CoAs by a long-chain acyl-CoA synthetase (LACS) (Andrews et al., 1983; Shockey et al., 2002; Schnurr et al., 2004; Lu et al., 2009). This esterification to CoA seems to prevent reabsorption into the plastid and increases solubility in the 	
    3	
    cytoplasm (Kunst et al., 2006). The resultant acyl-CoA substrates are elongated to chains 20 through 34 carbons long, via the cycle of four repetitive reactions performed by the fatty acid elongase (FAE) multi-enzyme complex located on the epidermal endoplasmic reticulum (ER) (von Wettstein-Knowles, 1982; Kunst et al., 2006; Samuels et al., 2008; Kunst and Samuels, 2009; Bernard et al., 2012). In each elongation cycle, a C2 moiety is added to the acyl-CoA substrate. The FAE is a heterotetramer that consists of four different enzymes. As in the fatty acid synthesis process, each cycle of fatty acid elongation is performed by four consecutive enzymatic reactions (Kunst and Samuels, 2003). First, the C16 or C18 fatty acyl-CoA substrate is condensed with a malonyl-CoA catalyzed by a β-hydroxy acyl-CoA synthase (KCS). Second, the β-keto group is reduced by a β-ketoacyl-CoA reductase (KCR) to produce β-hydroxy acylCoA. Third, it is dehydrated into trans-enoyl-CoA by β-hydroxyacyl-CoA dehydratase (HCD). Fourth, the enoyl-CoA is reduced by an enoyl-CoA reductase (ECR) to generate an acyl-CoA longer by two carbons.  The substrate specificity of each elongation reaction is thought to be determined by the condensing enzyme KCS (Millar and Kunst., 1997; Millar et al., 1999; Blacklock and Jaworski., 2006; Paul et al., 2006). A large family of twenty-one KCS-like sequences has been identified in the Arabidopsis genome (Blacklock and Jaworski, 2006; Joubes et al., 2008). Consistent with a role in wax synthesis, eight KCSs were found up-regulated in the Arabidopsis stem epidermis (Suh et al., 2005). Six of these KCSs (KCS1, KCS2, KCS10/FDH, KCS13/HIC, KCS20 and KCS6/CER6) have been studied in some detail and their role in wax precursor synthesis has been proposed (Gray et al., 2000; Yephremov et al., 1999; Todd et al., 1999; Millar et al., 1999; Pruitt et al., 2000; Lee et al., 2009). However, CER6 is the only wax-specific KCS characterized to date (Millar et al., 1999). CER6 is expressed specifically in the epidermis and the Arabidopsis cer6 mutant shows a decrease in wax compounds with C26 or longer chain lengths and 	
    4	
    simultaneous increase of C24 wax compounds, indicating that CER6 is involved in elongating C24 fatty acyl-CoAs (Todd et al., 1999; Millar et al., 1999). Furthermore, when CER6 was heterologously expressed in Saccharomyces cerevisiae, it produced fatty acids up to C28 in chain length (Tresch et al., 2012). Very recently, another yeast experiment has demonstrated that expressing CER6 together with an additional enzyme, CER2 (Negruk et al., 1996), produced C30 fatty acid (Haslam et al., 2012). This evidence indicates that elongation of C28 acyl-CoA to C30 acyl-CoA requires of both of these enzymes.  In contrast to KCSs, KCR, HCD and ECR are thought to have broad substrate chain length specificities (Beaudoin et al., 2009; Zheng et al., 2005; Bach et al., 2008). These genes, KCR, HCD and ECR, have been found and characterized in plants as well. Two maize genes GL8a and GL8b have been found to have overlapping expression patterns and function as KCR during VLCFA synthesis for wax production (Xu et al., 1997; Dietrich et al., 2005) A BLAST search using the yeast KCR sequence has identified two putative homologues in the Arabidopsis genome, both of which are highly expressed in the stem epidermis (Suh et al., 2005). One of these genes (At1g67730) rescued the growth phenotype of a KCR-defective yeast mutant (Beaudoin et al., 2002). The Arabidopsis CER10 gene encodes the ECR (Gable et al., 2004). Once again, evidence from yeast experiments showed that when CER10 was heterologously expressed in a yeast mutant lacking ECR activity, the wild-type phenotype of yeast was restored. Two Arabidopsis loci were identified as encoding for potential HCD activity (Bach et al., 2008). Complementation of the corresponding yeast mutant revealed that only PAS2 encodes a functional HCD in Arabidopsis (Bellec et al., 2002; Bach et al., 2008; Beaudoin et al., 2009). Indeed, in the last decade, major breakthroughs on VLCFA synthesis in yeast has paved the way for identifying the corresponding ECR, KCR and HCD enzymes in Arabidopsis (Kohlwein et al., 2001; Han et al., 2002; Beaudoin et al., 2002; Denic and Weissman., 2007). 	
    5	
    1.5	
  The	
  biosynthesis	
  of	
  wax	
  compounds	
    	
    It is widely believed that after fatty acid elongation, wax biosynthesis splits into the primary alcohol and alkane pathways. The former converts VLC-fatty acyl CoAs into primary alcohols and further into wax esters (Kunst and Samuels, 2003). Alternatively, the alkane pathway produces aldehydes, alkanes, secondary alcohols, and ketones (Cheesbrough and Kolattukudy, 1984; Schneider-Belhaddad and Kolattukudy, 2000). The two separate pathways are discussed independently in the following subsections (1.5.1 and 1.5.2).  1.5.1	
  Synthesis	
  of	
  primary	
  alcohols	
  and	
  wax	
  esters	
   Primary alcohol synthesis was shown biochemically to occur via a two-step reaction where VLCFAs are reduced to aldehydes and subsequently reduced to primary alcohols (Kolattukudy, 1980). Further biochemical studies in jojoba (Simmondsia chinensis) seeds and pea leaves, including functional expression of genes encoding alcohol-forming fatty acyl reductases (FAR) in yeast showed that alcohol synthesis from VLCFAs can be accomplished by a single enzyme, the potential aldehyde intermediate left bound to the FAR (Kolattukudy, 1971; Pollard et al., 1979; Vioque and Kolattukudy, 1997; Metz et al., 2000; Rowland and Domergue, 2012). Eight FAR-like genes were identified in the Arabidopsis genome, and one of them was identified as CER4 (Rowland et al., 2006). The Arabidopsis cer4 mutant shows a significant decrease in primary alcohols and wax esters, while the amounts of alkanes, secondary alcohols and ketones were not changed, indicating that CER4 has a role in the primary alcohol pathway (Jenks et al., 1995). While a mutation in CER4 drastically reduced the synthesis of C24 to C28 primary alcohols, C30 species were still detected in minor quantities, suggesting that an undiscovered FAR or a related functional enzyme may act in C30 primary alcohol synthesis (Rowland and Domergue, 2012). Although FAR activity of CER4 was confirmed in the yeast system where VLC-primary alcohols with C24 and C26 carbon chain lengths were produced, the activity was 	
    6	
    never demonstrated on C28 and C30 fatty acids (Rowland et al., 2006). Subcellular localization of green fluorescent protein GFP-tagged CER4 demonstrated its localization to the ER of epidermal cells in leaves and stems (Rowland et al., 2006). All these studies support CER4’s role as a FAR in cuticular primary alcohol synthesis.  The synthesized primary alcohols by the CER4 enzyme serve as substrates for wax ester production (Lai et al., 2007). The C16 acyl-CoA chain is the predominant acyl substrate for ester synthesis in Arabidopsis stem wax (Lai et al., 2007). Wax synthase (WS) enzymes catalyze the esterification of acyl-CoAs to primary alcohols in higher plants, mammals and prokaryotes (Lardizabal et al., 2000; Metz et al., 2000; Cheng and Russell, 2004; Stoveken et al., 2005). In the  Arabidopsis  genome,  eleven  genes  were  annotated  as  bi-functional  wax  synthase/diacylglycerol acyl transferase (WS/DGAT) genes (Waltermann et al., 2007). Analysis of one putative WS/DGAT encoding gene (WSD1) was shown to be involved in wax biosynthesis when up-regulated in the epidermis (Li et al., 2008). The corresponding wsd1 mutant had a severe reduction in wax esters and the WS activity was confirmed for C16 fatty acid and VLC-primary alcohols in yeast (Li et al., 2008). WSD1 was shown to localize to the ER in epidermal cells, consistent with the function in wax biosynthesis.  1.5.2	
  Synthesis	
  of	
  alkanes	
   VLC acyl-CoAs can be directed to the alkane-forming pathway to produce odd-numbered alkanes, secondary alcohols and ketones. In this subsection, the history of early work on alkaneformation is discussed, specifically the work leading to the simple two-step pathway hypothesis. Evidence for and against this hypothesis is also given.  	
    7	
    Kolattukudy and co-workers in the 1960s and 1970s used various plant species such as Brassica oleracea, Pisum sativum and Spinacia oleracea to do feeding experiments, showing that VLCFA precursors are taken up for the biosynthesis of alkanes and subsequently for secondary alcohols and ketones (Kolattukudy, 1965; Kolattukudy, 1968; Kolattukudy et al., 1972; Kolattukudy et al., 1971). These biochemical studies were later confirmed by molecular genetic studies with two Arabidopsis mutants, cer6 and cer10, that have mutations in elongating VLCFAs and showed significant reductions of longer acyl chains along with diminished levels of all the odd-numbered alkanes, secondary alcohols and ketones (Hooker et al., 2007; Hooker et al., 2002; Jenks et al., 1995; Rashotte et al., 2001; Zheng et al., 2005). Thus, both the biochemical and molecular genetic work shows that wax precursors are first elongated to VLCFA and then modified into alkanes, secondary alcohols and ketones.  In the 1980s, Kolattukudy et al. used isotope labeling and microsomal preparations from Pisum sativum to show that production of an alkane is accomplished by conversion of an aldehyde to an alkane and CO by a decarbonylation reaction (Cheesbrough and Kolattukudy, 1984). This reaction takes the even-numbered aldehyde and produces the odd-numbered alkanes, thereby the loss of one carbon. The reaction was suggested to be an enzymatic process since the conversion rates were affected by temperature, pH, time-course, substrate concentration, protein concentration and trypsin treatments. Thus, the enzyme proposed was a decarbonylase. Furthermore, Kolattukudy’s group reported that the enzymatic decarbonylation was inhibited by the presence of a metal ion-chelating agent, pointing to the importance of a metal ion in participating in the decarbonylation of octadecanal to heptadecane (Cheesbrough and Kolattukudy, 1984). In addition, they reported that an acyl-CoA reductase formed the aldehyde that serves as the substrate in the ensuing decarbonylation. Thus, the original two-step pathway reported by Kolattukudy and his group included the reduction and decarbonylation to yield 	
    8	
    alkanes. However, it should be noted that the in vitro characterization of the decarbonylase activity was carried out using octadecanal, an aldehyde with only 18 carbons and thus very different from the postulated physiological substrate with 30 carbons. Interestingly, the authors concluded the publication by stating that the mechanism cannot be understood without first solubilizing and characterizing the enzyme. Indeed, biochemical evidence on the enzyme is still lacking, possibly because the reductase and decarbonylase that are suggested to be encoded by CER3 and CER1 in Arabidopsis thaliana (see below), respectively, are both integral membrane proteins difficult to solubilize and study.  Two Arabidopsis genes, CER1 (Aarts et al., 1995) and CER3 (Kurata et al., 2003; Rowland et al., 2007) have been cloned and characterized to a certain degree (Aarts et al., 1995; Chen et al., 2003; Rowland et al., 2007; Xia et al., 2010; Bourdenx et al., 2011), and are thought to be involved in alkane formation. The first evidence came from chemical analyses of corresponding Arabidopsis mutant waxes. The stem wax of cer3 exhibited a severe reduction in all constituents of the alkane-forming pathway, while the amount of C30 primary alcohol was increased. The stem wax of cer1 mutants had a severe reduction in all alkanes, especially C29, and alkane metabolites; however, the level of C30 aldehyde increased slightly (Jenks et al., 1995). Furthermore, overexpression of CER1 in Arabidopsis led to a significant increase in alkanes in both stems as well as rosette leaves (Bourdenx et al., 2011). Consistent with a function in wax synthesis, CER1 and CER3 were found to be expressed in aerial organ epidermis and are upregulated in drought conditions precisely when wax production is especially required (Kurata et al., 2003; Kosma et al., 2009; Bourdenx et al., 2011). In addition, Goodwin et al (2005) have analyzed several double mutant lines including cer1cer3, and their data suggests that CER3 is upstream of CER1. However, the stem chemical analysis of cer1 did not show an elevation of C30 aldehyde, which greatly deviated from the previous results by Jenks et al (1995), casting 	
    9	
    doubts on the accuracy and reliability of both the single and double mutant data. Thus, a repeat of the chemical analysis of the double mutant is needed to confirm the sequence of CER1 and CER3 on the pathway. From the above-mentioned results it may be concluded that alkane formation occurs via CER3, an acyl-CoA reductase, converting acyl-CoAs to aldehydes and CER1 is the decarbonylase converting even-numbered aldehydes to odd-numbered alkanes.  Apart from Arabidopsis phenotypic evidence in favor of the two-step pathway hypothesis, direct evidence for involvement of CER1 and CER3 in the alkane-forming pathway came quite recently by heterologously reconstructing the plant alkane biosynthesis pathway in yeast (Bernard et al., 2012). Expression of CER1 or CER3 alone in a VLCFA-synthesizing yeast did not produce any change in the yeast lipid composition (Bernard et al., 2012). Co-expressing the two proteins in a yeast strain engineered to produce C30 fatty acid substrate, led to the synthesis of a new lipid identified as C29 alkane (Bernard et al., 2012). The amount of C29 alkane produced could be significantly increased by also expressing CYTB5-b in yeast, an electron transferring protein localized to the ER. This implies that a reducing system was required for alkane biosynthesis in Arabidopsis.  It must be noted that trans-membrane domains (Figure 3) have been predicted in the amino acid sequences of both CER1 and CER3. This is consistent with their subcellular localization to the ER. Furthermore, three histidine-rich motifs (Figure 4) have also been found in both proteins (Aarts et al., 1995; Chen et al., 2003; Kurata et al., 2003). To investigate the functions of the histidines in histidine-rich motifs of CER1 and CER3, one histidine of each histidine-rich motif was altered by site-directed mutagenesis to an alanine (Bernard et al., 2012). The reconstitution of alkane biosynthesis in yeast was unsuccessful when the mutated CER1 was expressed (Bernard et al., 2012). In addition, the mutated CER1 construct could not restore the Arabidopsis 	
    10	
    cer1 mutant phenotype. Interestingly, mutating the histidines in CER3 did not have any effect. Together, the results demonstrated that the histidine-rich motifs are indispensable (Shanklin et al., 1998; Shanklin et al., 2009) for the function of CER1 only (Bernard et al., 2012), and are not essential for CER3 (Bernard et al., 2012). To date, the current status of research on the function of CER1 and CER3 is incomplete, since more biochemical evidence is still required to (1) identify the substrate, and (2) the function of these enzymes.  Despite the evidence given in support of the two-step pathway hypothesis, there is substantial evidence against it. The CER1 and CER3 protein sequences share 35% of amino acid similarity and both enzymes are integral membrane proteins with similar domain structures (Figure 4). Their N-terminal ends show primary sequence similarity with fatty acid desaturases (NCBI Conserved Domain Search) and the C-terminal ends with short chain dehydrogenases (SDRs). However, it should be noted that the two domains in the two enzymes have been annotated as having oxidase functions, which does not correspond with the earlier mentioned sequence of reductions and C-C bond cleavage.  Further evidence casting doubt on the simple two-step pathway stems from two more genes potentially being involved in the process. First, CER8 was recently found to be identical to LACS1, one of nine Arabidopsis long-chain acyl CoA synthetases that activate acyl chains (Lu et al., 2009). The Arabidopsis cer8 mutant has reduced amounts of alkanes together with increased levels of C30 free fatty acids (Lu et al., 2009). Second, a patent was recently published claiming that Susceptible to Coronatine-Deficient Pst DC3118-2 (SCD2) can convert aldehydes to alkanes (Zeng and He, 2010). The scd2 mutant was found to accumulate aldehydes while lacking alkanes, and this phenotype was restricted to leaves rather than stems. Thus, it is suspected that SCD2 performs the same function in leaves as CER1 does in stems (Zeng and He, 	
    11	
    2010). However, SCD2 is annotated as a thioesterase, and an apparent function on the simple pathway cannot be assigned. To provide more evidence for its involvement in alkane-formation, full characterization of SCD2 is needed.  Recently, work has been done on cyanobacterial alkane biosynthesis where evidence was presented for a classical sequence of reactions where the acyl CoA reductase forms aldehydes from acyl CoAs and a decarbonylase forms alkanes from aldehydes (Schirmer et al., 2010). However, the two cyanobacterial proteins showed no significant sequence similarity with either CER1 or CER3, either as a whole or on the domain level, indicating that alkane formation pathway must be more or less different in plants.  1.5.3	
  Synthesis	
  of	
  secondary	
  alcohols	
  and	
  ketones	
   Alkanes serve as substrates for the production of secondary alcohols and ketones. The stem wax of Arabidopsis contains up to 30% nonacosan-15-ol and nonacosan-14-ol, as well as small amounts of other secondary alcohol isomers. Nonacosan-15-one is the predominant ketone in Arabidopsis stem wax. A cytochrome P450-dependent enzyme called mid-chain alkane hydroxylase (MAH1) was identified in a reverse genetic approach in Arabidopsis (Greer et al., 2007). The corresponding mah1 mutant had close to total reduction of secondary alcohols and ketones. Interestingly, lines ectopically expressing MAH1 in leaves had an increase in secondary alcohols and ketones, where in wild type leaves they are found only in trace amounts. Subcellular localization of MAH1 to the ER was in line with its role in wax biosynthesis.  1.6	
  Research	
  questions	
  and	
  objectives	
   The current Master’s work was focused on studying the alkane-forming pathway in Arabidopsis, the least understood branch of wax biosynthesis. The research in this thesis was aimed at 	
    12	
    addressing the following over-arching question: Is alkane-formation governed by a two-step pathway? Is there additional evidence against it?  In order to explore further the involvement of an additional protein SCD2 in the Arabidopsis alkane biosynthesis pathway, a complete characterization was undertaken in Chapter 3. In particular, the following questions were addressed: 1) Are the amounts of alkanes reduced and aldehydes increased in leaves of scd2-1 and scd2-2 single mutant lines? 2) Is SCD2 solely responsible for the observed mutant chemical phenotype? 3) What is the organ-specific expression pattern of SCD2? 4) What is the tissue-specific expression pattern of SCD2? 5) What is the subcellular localization of SCD2? In Chapter 4, CER1 and CER3 were studied in more detail by generating the Arabidopsis cer1cer3 double mutant to ascertain the existence of epistatic interactions. The following questions were addressed: 6) What is the order of CER3 and CER1 in the alkane-forming pathway? Is the chemical phenotype the same or different in stems and leaves of cer1cer3 double mutants and their respective single parents? 7) Does the double mutant display sterility? And to what degree does it do so compared to the single mutant parents? A complete characterization of the double mutant was carried out in comparison to the wild type and the respective single mutants. This included analyzing the wax chemical phenotype both qualitatively by employing scanning electron microscopy (SEM) and quantitatively with gas chromatography-flame ionization detection (GC-FID) and then gas chromatography-mass spectrometry (GC-MS).  	
    13	
    Finally, additional cer1cer4 and cer3cer4 double mutants were generated to study the branchpoint of the wax biosynthetic pathway.  I used molecular genetics, confocal microscopy and analytical chemistry techniques to address the above-mentioned questions.  	
    14	
    	
   Figure 1 Scheme showing a cross-section of a plant cuticle and epidermal cells. Figure modified after Jetter et al (2000) and Jeffree (1996). 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    15	
    	
   Figure 2 Wax Biosynthetic Pathway in Arabidopsis thaliana stems. CER, eceriferum; WSD, wax synthase/diacylglycerol acyltransferase; MAH, mid-chain alkane hydroxylase. Figure modified from Samuels et al (2008). 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    16	
    	
    	
   	
   Figure 3 Trans-membrane prediction results of CER1 and CER3 presented by the TMHMM program. 	
   	
   	
   	
   	
   	
    17	
    	
   	
   Figure 4 Scheme of CER1 and CER3 proteins with desaturase and dehydrogenase (SDR) domains illustrated. Histidine-rich motifs are represented with orange boxes. The following scheme of domains was predicted using the hydropathy results of the TMHMM program.  	
    18	
    Chapter	
  2	
  Materials	
  and	
  methods	
   2.1	
  Plant	
  materials	
  and	
  growth	
  conditions	
   Single mutant lines SALK_038693 (cer4) (Col-0 ecotype), cer1-2 SALK_014839 (Col-0 ecotype) and cer3-6 (yre Col-0 ecotype) were used to cross and generate the following double mutant lines: cer1-2cer3-6, cer1-2cer4, and cer3-6cer4. Mutant seeds of cer4 were obtained from the ABRC (www.arabidopsis.org). The mutant seeds of cer1-2 and cer3-6 alleles were a kind gift from Dr. Owen Rowland (Carleton University). The mutant seeds of scd2-1 and scd2-2 were a kind gift from Dr. Weiqing Zeng. Seeds were stratified for 2 to 3 days at 4°C and subsequently germinated on AT-agar plates without antibiotics at 20°C under continuous light (100 µE m-2 s-1 of photosynthetically active radiation). The 14 day-old seedlings were then transplanted onto soil (Sunshine Mix 5; SunGro) and watered every three days in the growth chamber at 22°C under long-day conditions of a 16-h-light/8-h-dark cycle.  2.2	
  Wax	
  analyses	
   2.2.1	
  Wax	
  extraction	
  and	
  derivatization	
   Five week-old Arabidopsis thaliana plants of various genotypes were used to determine wax composition. Stems and rosette leaves were first collected and photographed with a digital camera for area analysis using the Image J program. Subsequently, samples were submerged in chloroform for 30 seconds to remove epi- and intracuticular wax. The solvent was removed from the wax solutions under a gentle stream of nitrogen gas. Wax was then derivatized by addition of (BSTFA) N,O-bis(trimethylsilyl) trifluoroacetamide and pyridine at 80°C for 40 min. The solvents were evaporated under a gentle stream of nitrogen gas and chloroform was added to the samples before quantitative analyses were performed by gas chromatography-flame ionization detection (GC-FID) and identification by gas chromatography-mass spectrometry (GC-MS). 	
    19	
    2.2.2	
  Chemical	
  analyses	
  using	
  GC-­‐FID	
  and	
  GC-­‐MS	
   Wax was analyzed using capillary GC (5890N, Agilent, Avondale, Pa; column 30 m HP-1, 0.32 mm i.d., df= 0.1 µm, Agilent) using temperature-programmed on-column injection at 50°C, oven for 2 min at 50°C, then raised by 40°C/min to 200°C, held for 2 min at 200°C, then raised by 3°C/min to 320°C, and finally held at 320°C for 30 min. Subsequent qualitative analysis was performed by GC connected to a mass spectrometric detector (5973N, Agilent) with helium gas inlet pressure programmed to 1.4 ml/min. Each wax constituent was identified first from characteristic fragmentation peaks using mass spectrometry, then matched to the corresponding GC-FID peaks. Peaks were integrated to determine the area under each of the peaks. The quantity (µg) was established by comparison with a defined amount of C24 alkane, the internal standard added to the wax extracts. Wax loads were calculated by comparing the GC-FID peak areas against the internal C24 alkane standard and dividing by the calculated surface area of each sample. The quantitative data are presented as means of parallel experiments with standard deviation (SD).  2.3	
  Genomic	
  DNA	
  extraction,	
  RNA	
  isolation,	
  cDNA	
  synthesis	
  and	
  reverse	
  transcription	
   2.3.1	
  Genomic	
  DNA	
  extraction	
  for	
  PCR	
  genotyping	
   Prior to PCR genotyping of each of the single and double mutant lines, genomic DNA was isolated for each sample that was screened. One true leaf was excised and submerged in 50 µl of extraction buffer (Sigma) and incubated for 10 min at 95°C. To each sample, 50 µl of dilution buffer (Sigma) was immediately added, vortexed and stored at -20°C until further use. Genespecific primers were used to amplify the wild-type locus. Lba1 primer was used to check for the presence of the T-DNA.  	
    20	
    2.3.2	
  RNA	
  and	
  cDNA	
  preparation,	
  quantitative	
  RT-­‐PCR	
  conditions	
  and	
  analysis	
   RNA was extracted from Arabidopsis Col-0 leaf tissue with RNeasy Plant mini kit (Qiagen) using liquid nitrogen and pestle and mortar. Purified RNA was treated with DNase I using the “DNase I kit” (Qiagen). First-strand cDNA was prepared from 1 µg of total RNA with the Superscript RT III kit (Invitrogen) and oligo(dT)18 primers according to the manufacturer’s instructions. A 0.8 µl aliquot of the total reaction volume (20 µl) was used as template in realtime reverse transcriptase (RT)-mediated PCR amplification reaction. Gene specific primers were used in the PCR amplification. PCR efficiency ranged from 95 to 100%. All samples were assayed in triplicate wells, and the PCR reaction was performed twice for each sample. Real-time PCR was performed using the iCycler  TM  (Bio-Rad, Hercules, CA, USA). Samples were  amplified in 20 µl reactions containing 1x SYBR Green Master Mix (Bio-Rad) and 100 mM of each gene-specific primer. The thermal conditions for SCD2 amplification were: 1 cycle at 95°C for 3 min and 30 sec followed by 40 cycles at 95°C for 30 sec, and at 60°C for 30 sec annealing. Data acquisition and analysis was performed using the iCycler TM IQ software (version 3.0a, BioRad) for each run of the real-time RT-PCR. Threshold cycles (CT) were adjusted and CT values for each housekeeping gene (ACT2, EF-1α, PP2A, UBQ10, and elF-4A-1) amplified on each 96 well PCR plate in parallel were subtracted from those of gene of interest to generate the normalized ΔCT values. The ΔΔCT values were generated by subtracting the CT values of an arbitrary calibrator (roots) from the ΔCT values. The final relative expression levels were calculated by the 2-ΔΔCT method.  2.4	
  Scanning	
  electron	
  microscopy	
  (SEM)	
  	
   Segments isolated from the apical part of the stem (1 cm) were mounted onto SEM stubs, air dried at room temperature overnight, and coated the next morning with gold particles in a 	
    21	
    SEMPrep2 sputter coater (Nanotech). The coated samples were viewed and imaged with a Hitachi S4700 field emission SEM with an accelerating voltage of 1 kV and a distance of 12 mm.  2.5	
  Cloning	
  and	
  subcellular	
  localization	
  of	
  SCD2	
  in	
  Arabidopsis	
  thaliana	
  and	
  Nicotiana	
   benthamiana	
   	
    To investigate the subcellular localization of SCD2, the coding sequence of SCD2 was fused to green fluorescent protein (GFP). Total RNA was extracted from wild-type Col-0 Arabidopsis rosette leaf by grinding the tissue in liquid nitrogen with pestle and mortar. RNA was extracted using the RNeasy Plant mini kit (Qiagen). Purified RNA was treated with DNase I using the “DNase I kit” (Qiagen). First-strand cDNA was prepared from 1µg of total RNA with the Superscript RT III kit (Invitrogen) and oligo(dT)18 primers according to the manufacturer’s instructions. PCR was performed to amplify the SCD2 cDNA fragment by using High Fidelity DNA Polymerase (New England Biolabs) with 1 µl of cDNA. Primers used for PCR amplification were gene-specific forward primer SCD2 N-GFP (5’- GGG GAC AAGTTT GTA CAA AAA AGC AGG CTT CAT GGGTGT GGT AGA AGA AGCT -3’) and reverse primer SCD2 N-GFP (5’- GGG GAC CACTTT GTA CAA GAA AGCTGG GTC TCA CAT AGC AAT GTC ATT ACG AAT GTG AC -3’) to generate the N-terminal fusion with GFP. To construct the C-terminal fusion with GFP, gene-specific forward primer SCD2 C-GFP (5’- GGG GAC AAGTTT GTA CAA AAA AGC AGG CTT CAT GGGTGT GGT AGA AGA AGCT -3’) and reverse primer SCD2 C-GFP (5’- GGG GAC CACTTT GTA CAA GAA AGCTGG GTC CAT AGC AAT GTC ATT ACG AAT GTG AC -3’) were used. The PCR thermal conditions were as follows: initial denaturing at 96°C for 1 minute and 30 sec, and 35 cycles of 57°C annealing for 20 sec and 72°C extension for 40 sec. First, the PCR-amplified SCD2 cDNA 	
    22	
    fragment of 816 bp was cloned into pDONR207 entry vector (Invitrogen) using BP Clonase II enzyme (Invitrogen) via the Gateway Mediated BP reaction and transformed into TOP10 chemically competent E.coli cells. Colonies were selected on Luria-Bertani (LB) solid medium containing 50 µg/ml hygromycin (w/v). Resistant colonies were inoculated in liquid LB medium supplemented with hygromycin (50 µg/ml) overnight for subsequent plasmid extraction. Plasmids were sequenced to verify that no mutations were introduced during the PCR amplification reaction. pDONR207 plasmids harboring the SCD2 cDNA destined for N- and Cterminal GFP fusion were subjected to the Gateway LR reaction using the LR Clonase II reaction mix (Invitrogen) to generate the expression vectors pGWB6-p35S:GFP-SCD2 (N-sGFP) and pGWB5-p35S:SCD2-GFP (C-sGFP) under the control of the constitutive cauliflower mosaic virus 35S promoter. Agrobacterium tumefaciens strain GV3101 (pMP90) was transformed with the resulting GFP fusion constructs by the heat-shock method (Sparkes et al., 2006). Agrobacteria with the plasmids pGWB6-p35S:GFP-SCD2 (N-sGFP) and pGWB5-p35S:SCD2GFP (C-sGFP) were plated onto LB medium supplemented with kanamycin (50 µg/ml), hygromycin (50 µg/ml), rifampicin (100 µg/ml) and gentamycin (50 µg/ml) and grown for 2-3 days at 28°C. Agrobacterium carrying p19 was grown at 28°C in LB medium containing kanamycin (50 µg/ml) and rifampicin (100 µg/ml). Each Agrobacterium tumefaciens colony was subsequently screened for presence of the plasmid by PCR. Transformation of scd-1 Arabidopsis plants was performed using the floral-dip method (Clough and Bent, 1998) and T1 transformants were selected on AT-agar medium containing 50 µg/ml of hygromycin (w/v). More than 100 resistant transformants were recovered for each transgenic line.  	
    23	
    2.5.1	
  Transient	
  expression	
  of	
  SCD2	
  in	
  Nicotiana	
  benthamiana	
   The two constructs, pGWB5-p35S:SCD2-GFP and pGWB6-p35S:GFP-SCD2, and an ERspecific control marker line p35S:HDEL-RFP were grown up from an Agrobacterium tumefaciens glycerol stock. Both the N. benthamiana seeds and the p35S:HDEL-RFP marker plasmid were generously provided by Dr. Mathias Schuetz (Samuels lab, Department of Botany, University of British Columbia). Four ml of LB liquid medium supplemented with hygromycin, kanamycin, gentamycin, and rifampicilin at 50 µg/ml each was grown overnight. The next day, Agrobacterium tumefaciens cells were centrifuged, washed with dH2O, and resuspended in 4 ml of dH2O. Leaves of three-week-old N. benthamiana plants were infiltrated with Agrobacterium suspension by applying a sterile syringe lacking a needle onto the underside of the leaf. The ERspecific control marker line p35S:HDEL-RFP was co-infiltrated with each construct. Infiltrated tobacco plants were returned to the growth chamber, and after three days the infiltrated leaves were checked for expression using a confocal microscope and imaged (BioImaging Facility, Department of Botany, University of British Columbia).  2.5.2	
  Light	
  microscopy	
  and	
  laser	
  scanning	
  confocal	
  microscopy	
   Transiently expressing N. benthamiana leaf tissues and stably-transformed Arabidopsis root tissues with the two constructs, pGWB5-p35S:SCD2-GFP and pGWB6-p35S:GFP-SCD2, were prepared and analyzed for GFP signal. N. benthamiana leaf tissues transiently expressing two constructs pGWB5-p35S:SCD2-GFP and pGWB6-p35S:GFP-SCD2 with the p35S:HDEL-RFP were analyzed for co-localization using an Olympus multi-photon confocal microscope (BioImaging Facility, Department of Botany, University of British Columbia) with an excitation of 488 nm for GFP and 543 nm for RFP, respectively. Images were processed using Fiji and Volocity softwares.  	
    24	
    2.5.3	
  Promoter-­‐GUS	
  cloning	
  and	
  histochemical	
  staining	
   To clone the promoter region of SCD2, wild-type Arabidopsis leaf gDNA was first extracted using DNAzol reagent (Invitrogen). A 1018 bp promoter region immediately upstream of the ATG start codon of SCD2 was amplified by PCR from the extracted gDNA. The primers used for amplification were gene-specific forward primer “Sequence-promoter SCD2 GUS” 5’- GGG GAC AAGTTT GTA CAA AAA AGC AGG CTT CTT CAC GAC CAGTAT GGTTTA CTC A -3’ and gene-specific reverse primer “Sequence-promoter SCD2 GUS” 5’- GGG GGA CCA CTT TGT ACA AGA AAG CTG GGT CCATCT CTC TAA AGA AGATTC TTC TCT GG -3’. The primers contain Gateway-specific sequences for recombination. The PCR product was subjected to a BP recombination reaction (Gateway) using BP Clonase II enzyme (Invitrogen) and cloned into the pDONR207 donor vector (Invitrogen). The resulting construct pDONR207/pSCD2 was first sequenced to check for possible mutations introduced during the PCR reaction and was recombined with the LR sites of pMDC162 to generate pMDC162/pSCD2:GUS using LR Clonase II enzyme mix (Invitrogen) in a Gateway LR reaction. The resulting expression vector pMDC162/ pSCD2:GUS was transformed into Agrobacterium tumefaciens strain GV3101 (pMP90) using the heat-shock method. Arabidopsis wild-type Col-0 plants were transformed using the floral-dip method (Clough and Bent, 1998) and transformants were selected on AT-medium containing 50 µg/ml hygromycin (w/v). More than 100 resistant T1 transformants were recovered.  GUS staining solution consisted of 50 mM sodium phosphate, pH 7.0, 2 mM potassium ferricyanide, 2 mM potassium ferrocyanide, 0.1% (v/v) Triton X-100, and 2 mg/ml 5-bromo-4chloro-3-indolyl-β-D-glucuronide (X-Gluc). Tissues submerged in GUS staining solution with X-Gluc were vacuum-infiltrated for 30 min at 5 min-intervals. Root and aerial tissues were incubated at 37°C for 1 h or overnight. The reaction was stopped by removing the staining 	
    25	
    solution and adding 100% EtOH. Samples were cleared of chlorophyll with serial additions of 70% EtOH and subsequently stored in 90% acetone. Tissues were visualized using a dissecting light microscope (Sack lab, Department of Botany, University of British Columbia).  	
    26	
    Chapter	
  3	
  Molecular	
  characterization	
  of	
  SCD2	
  	
   3.1	
  Introduction	
   A recent patent (Zeng and He, 2010) described an Arabidopsis gene, SCD2 (At4g37470), whose corresponding protein may be an enzyme catalyzing the reduction of aldehydes to alkanes. The authors partially characterized one mutant allele of SCD2 (scd2-1), a point mutant having a single nucleotide deletion that causes an early stop codon. The authors presented a second mutant allele of SCD2, a Ds element insertion scd2-2 (CS100282) that was genotyped, but chemical analyses of cuticular wax were not reported. These two alleles are in different genetic backgrounds. The authors performed chemical analysis on the homozygous lines of scd2-1 and reported (Figure 5) (Zeng and He, 2010) an increase in C30, C32 and C34 aldehyde levels and a decrease in the C29, C31 and C33 alkane levels in leaves, suggesting that SCD2 is involved in alkane formation in leaves. Furthermore, stem wax (Figure 5) as well as leaf and stem cutin (Figure 6) were unaltered compared to the Col-7 wild type. The evidence presented in the patent (Zeng and He, 2010) only contained wax and cutin chemical analyses for scd2-1 and not for the second mutant allele scd2-2.  Wax analysis was performed on several transgenic plants expressing the SCD2 cDNA under the control of the native promoter and that of the constitutive cauliflower mosaic virus 35S promoter in the scd2-1 background, which showed that alkane levels were restored to wild type levels. Unfortunately, the quality of the wax data from the transgenics is questionable, since no evidence was presented that these lines were genotyped for the presence of the transgene. The authors also reported that transgenic plants overexpressing the SCD2 cDNA under control of the native promoter exhibited a drought resistant phenotype in leaves compared to the wild type (Zeng and  	
    27	
    He, 2010). Surprisingly, the scd2-1 mutants were susceptible to drought even though the total leaf wax load did not differ from wild type (Zeng and He, 2010).  Thus, independent confirmation of the wax phenotype in a second allele, if possible, combined with high-quality wax data on complementation lines, is necessary to prove that SCD2 is involved in alkane formation. This could be further corroborated with experiments testing for SCD2’s organ-/tissue-expression and subcellular localization, by promoter-GUS analysis or GFP, respectively.  The same Arabidopsis gene, At4g37470, has been characterized under a different name HTL (hyposensitive to light) and was shown to be implicated in photomorphogenesis. The promoter activity was found in the vasculature (Sun and Ni, 2010). Two htl-1 and htl-2 alleles both displayed a long hypocotyl phenotype under red, far-red and blue light, whereas overexpression of HTL caused a short hypocotyl phenotype under similar light conditions. The mutants also showed other photomorphogenic defects such as elongated petioles, retarded cotyledon and leaf expansion. HTL was strongly localized to the nucleus and weakly to the cytosol. This report did not provide a clear role for HTL and the characterization was incomplete. In a different photomorphogenesis study, At4g37470 was described as an alpha/beta hydrolase KAI2 (karrikin insensitive) (Waters and Smith, 2012) necessary for responses to karrikins, essential for normal light-dependent seedling development. Wax characterization in those two papers was not reported. No other study published, apart from the 2010 patent, has studied SCD2 as having any role in wax biosynthesis.  The current work was focused on addressing the following question: Is SCD2 indeed involved in alkane formation, as stated in the patent? If it is, then SCD2 will be an additional gene product 	
    28	
    that challenges the current two-step model of the alkane-forming branch of wax biosynthesis in Arabidopsis (discussed in 1.5.2). If so, what is the spatial expression pattern of SCD2 and does it correlate with previously reported wax-related genes? To answer these main questions, the following sub-questions were addressed experimentally: 1) Are the leaf alkane levels reduced and aldehyde levels increased in scd2-1 and scd2-2? 2) Can leaf wax be restored by transgene complementation? 3) What is the organ-specific expression pattern of SCD2 in Arabidopsis? 4) What is the tissue-specific expression pattern of SCD2 in Arabidopsis? 5) What is the subcellular localization of SCD2 in both Nicotiana benthamiana and Arabidopsis?  The transgene construct under control of the native SCD2 promoter was cloned and transformed into scd2-1 by Dr. Weiqing Zeng who provided the seeds. I generated the p35S:SCD2-GFP and pSCD2:GUS. I performed all subsequent steps, including transformation into scd2-1 mutant plants and Col-0, respectively, and analysis for not only subcellular localization in Nicotiana benthamiana and Arabidopsis, but also for all transgenic lines. The only materials not generated by me were scd2-1, scd2-2, and pSCD2:SCD2 in scd2-1.  3.2	
  Results	
   Both single mutant lines, scd2-1 and scd2-2, exhibited no obvious deviations in morphology from wild type. A glossy stem phenotype associated with several eceriferum (cer) mutants (Koornneef et al., 1989; Hannoufa et al., 1993, McNevin et al., 1993) was not observed. Furthermore, no indication of post-genital organ fusions was observed that is often linked with a compromised cuticle. Chemical analysis of scd2-1 and scd2-2 stem and leaf wax was necessary to study the subtle alterations in their cuticles. 	
    29	
    In order to obtain stem and leaf wax profiles of scd2-1 and scd2-2, cuticular waxes from both organs were extracted with chloroform from four-week-old plants of Col-7, scd2-1, Ler and scd2-2. The amounts of each of individual compounds belonging to free fatty acids, primary alcohols, aldehydes, alkanes, secondary alcohols, ketones, triterpenoids, and other unidentified compounds were quantified by gas chromatography with flame ionization detector (GC-FID) and identified by their characteristic mass spectra using gas chromatography and mass spectrometry (GC-MS). Additionally, two representative complementation lines pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 were also examined for stem and leaf wax profiles.  3.2.1	
  Total	
  stem	
  wax	
   The total stem wax loads were 2401 ± 65 µg/dm2 for Col-7, 2389 ± 70 µg/dm2 for scd2-1, 2413 ± 57 µg/dm2 for Ler, and 2397 ± 62 µg/dm2 for scd2-2. All four wax mixtures contained the same compound classes in very similar quantities (Figure 7). The most abundant compound class was straight-chain alkanes amounting to 35% of the total wax. Approximately 25% of the wax was comprised of the ketone nonacosan-15-one, and 10% were the secondary alcohols nonacosan-14-ol and nonacosan-15-ol. The wax also contained primary alcohols (10%), esters (6%), aldehydes (3.5%), fatty acids (1%) and triterpenoids (0.8%). The rest of the wax (10%) was not identifiable.  The chain length distribution within each compound class of Col-7, scd2-1, Ler, scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 stem wax was further investigated (Figure 8). In all four lines, alkanes ranged from C27 to C33 and were dominanted by C29 (91%). Although odd-numbered alkanes were found to be most abundant, even-numbered alkanes C28 and C30 were also present, albeit in trace (<0.7%) quantities. All other compound classes were dominated by even-numbered chain lengths. Primary alcohols ranged from C24 to 	
    30	
    C32 and were dominated by C28. Equal ratios of C26 and C30 were found. Fatty acids were dominated by C28 compounds and aldehydes contained equal ratios of C28 and C30 compounds. Esters ranged from C40 to C50, and the most abundant ester had 44 carbons, containing the most prevalent alcohol C28 and C16 acid. The triterpenoids mainly contained β-amyrin and trinorlupeol, together at (<1%) of the total wax.  The total wax load on stems of pSCD2:SCD2 and p35S:SCD2-GFP in scd2-1, did not differ from that of wild type (Figure 7). The compound class compositions did not differ from the wild type either, in both absolute and relative quantities (Figure 7). Furthermore, the chain length distribution of compounds within their respective classes did not display any observable differences between the transgenic lines and the WT (Figure 8).  3.2.2	
  Total	
  leaf	
  wax	
   The total wax load for Col-7, scd2-1, Ler, scd2-2, pSCD2:SCD2 and p35S:SCD2-GFP was 119.5 ± 7.1 µg/dm2, 112 ± 6.4 µg/dm2, 110 ± 6.2 µg/dm2, 92.3 ± 6.6 µg/dm2, 120 ± 7.0 µg/dm2 and 119 ± 7.4 µg/dm2, respectively (Figure 9 and Table 1).  Only four compound classes were identified within the total leaf wax mixture (Table 1). Alkanes represented the most abundant class with approximately 46% (55.4 ± 3.8 µg/dm2) of the Col-7 wax. Primary alcohols (26.7 ± 1.8 µg/dm2) and aldehydes (13.3 ± 0.7 µg/dm2) contributed approximately 20% and 10% respectively each to the total wax load. Fatty acids (3.6 ± 0.1 µg/dm2) were the least abundant class. The rest (23.0 ± 1.1 µg/dm2) of the wax was not identified.  	
    31	
    Compared to the Col-7 compound class loads, the scd2-1 mutant had no difference in fatty acids and primary alcohols (Table 1). However, the aldehydes (30.6 ± 2.1 µg/dm2) of scd-1 were 231% of Col-7, while the alkanes (35.9 ± 2.8 µg/dm2) were only 64% of Col-7. The scd2-2 mutant displayed no difference in fatty acids and primary alcohols compared to its respective wild-type background Ler, similar to Col-7 and scd2-1. Aldehyde levels (28.1± 2.2 µg/dm2) were 226% of Ler and the alkane levels (14.5 ± 1.1 µg/dm2) were 31% of Ler, a more drastic change than was seen for Col-7 and scd2-1 (Table 1).  The two representative complementation lines pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1, had similar loads of all compound classes to Col-7 (Table 1).  The chain length distribution within each compound class of Col-7 leaves was further analyzed (Figure 10). Alkane chain lengths ranged from 27 to 37 carbons and were dominated by C31 and C29 at 49% and 27%, respectively. Similar to Arabidopsis wild-type stems, alkanes were dominated by odd-numbered chain lengths, however, even-numbered alkanes C30 and C32 were also identified in limited quantities (2%). The rest of the compound classes were composed primarily of even-numbered compounds with minor contributions from odd-numbered carbon compounds. The primary alcohols consisted of straight-chain and branched compounds. The straight-chain primary alcohols ranged from C26 to C34 and were dominated by C28 (6.3 ± 0.9 µg/dm2) and C26 (4.1 ± 1.1 µg/dm2) chain lengths. The branched primary alcohols ranged from C30 to C34 and were dominated by C32 (6.2 ± 1.1 µg/dm2) and C30 (3.5 ± 0.8 µg/dm2) chain lengths. Free fatty acids exhibited a chain length distribution from C24 to C34. The predominating free fatty acid was C26. Aldehydes also had a wide chain length range. The C32 aldehyde was the most abundant in its class. 	
    32	
    Compared to Col-7, the leaf wax of the scd2-1 single mutant contained the same chain lengths in all the compound classes (Figure 10). The relative amounts of chain lengths within fatty acids and primary alcohols were unchanged. In sharp contrast, scd2-1 had a dramatic rise in C30, C32, and C34 aldehyde levels, to 560%, 176% and 182% of Col-7, respectively. Alkanes declined dramatically, especially C29, C31, and C33 having 57%, 61% and 55% of Col-7, respectively (Figure 10).  Similar to changes from Col-7 to scd2-1, scd2-2 differed from Ler in aldehydes and alkanes (Figure 10). Specifically, aldehydes of chain lengths C30, C32, and C34 were 545%, 179% and 186% of Ler, respectively. Alkanes of chain lengths C29, C31 and C33 were 23%, 20.8% and 25% of Ler, respectively. The two representative complementation lines pSCD2:SCD2 and p35S:SCD2-GFP in scd2-1 exhibited no changes in any of the compound class chain length loads compared to Col-7 (Figure 11).  3.2.3	
  Organ-­‐specific	
  expression	
  of	
  SCD2	
   To investigate where SCD2 is expressed and whether the organ expression correlates with the leaf-specific chemical phenotype, I analyzed SCD2 transcript levels in Col-0 using quantitative RT-PCR rather than semi-quantitative RT-PCR. In many cases, when genes are strongly expressed or exhibit highly different expression levels in various organs, a semi-quantitative RTPCR is sufficient to visually ascertain differences in band saturation on an agarose gel. However, when a gene is expressed weakly, the subtle increase and/or decrease in expression is ideally measured by qRT-PCR .  Quantitative RT-PCR analyses were performed on four-week old Arabidopsis plants. The SCD2 expression was assayed in various organs and was compared against the house-keeping gene β	
    33	
    actin. The expression range of SCD2 in all organs was much lower than that of β-actin (Figure 12). A clear difference in expression level between various organs was observed. The highest abundance of the SCD2 transcript was in cauline (0.246 ± 0.011) and rosette leaves (0.174 ± 0.03), followed by siliques (0.136 ± 0.009). The stem as a whole exhibited lower level of expression (0.134) than did either rosette or cauline leaves. Modest expression levels were seen in the bottom (0.05 ± 0.01) and mid-regions (0.068 ± 0.002) of the stem, whereas the top of the stem had very low expression (0.016 ± 0.001). Flowers had expression levels similar to the stem (0.059 ± 0.012), and particularly low transcript levels were detected in the roots (0.021 ± 0.003).  3.2.4	
  Tissue-­‐specific	
  expression	
  of	
  SCD2	
   To further investigate the spatial expression pattern of SCD2, I explored the tissue expression profile of the gene. A promoter region 1018 bp upstream of the start codon of SCD2 was fused to the β-glucuronidase (GUS) reporter gene and the construct used to transform Arabidopsis. GUS activity was tested in two-week-old T2 pSCD2:GUS transgenic seedlings.  GUS activity was detected in the vasculature of cotyledons and more weakly in the first true leaves (Figure 13). GUS expression was also found in the phloem of the vasculature of leaves and not the xylem. In addition, the GUS reporter gene was expressed in the vasculature along the root, except for the root tip, and in the hypocotyl (Figure 13). Overall, SCD2 expression seems to be specific to the phloem of all the seedling organs, including the shoot, cotyledons, true leaves and roots. Using higher magnification, the phloem-specific expression was confirmed in the midvein of the true leaf (Figure 13L), indicating possible unknown functions for SCD2 in the vasculature.  	
    34	
    3.2.5	
  Subcellular	
  localization	
  of	
  SCD2	
   To further test whether SCD2 may be directly involved in alkane formation, I investigated the subcellular localization of this protein. In preparation for this, the amino acid sequence of SCD2 was first used for in silico analysis of possible trans-membrane domains using the TMHMM online program. SCD2 was predicted to lack trans-membrane domains (Figure 14), as well as a nuclear localization signal or other signaling peptides.  To verify the computational prediction, the subcellular localization of SCD2 was investigated with two constructs where SCD2 was fused with the green fluorescent protein (GFP) at the N- or C-terminus, and expressed under the control of the constitutive cauliflower mosaic virus 35S promoter. Both constructs were used for subcellular localization initially through Nicotiana benthamiana transient expression (Section 3.2.6), and then employing stable transformants of Arabidopsis (Section 3.2.7). GFP fusion with the native promoter was being performed by Dr. Weiqing Zeng independently.  3.2.6	
  Transient	
  expression	
  of	
  SCD2	
  in	
  Nicotiana	
  benthamiana	
   Two week-old N. benthamiana leaves were used to co-infiltrate two constructs: p35S:SCD2-GFP and p35S:HDEL-RFP into the underside of the leaf. The GFP fluorescence was detected in transgenics harboring N- and C-terminal fusion with GFP, indicating that the fusion proteins were not silenced and folded properly, and that they could be used for localization studies. However, since the C-terminal fusion produced stronger GFP signal, only the results for the Cterminal fusion p35S:SCD2-GFP are reported here (Figures 15 and 16). The pavement cells showed a reticulate pattern of fluorescence that is indicative of an endoplasmic reticulum (ER) membrane (Figure 15A). The pavement cells expressing the GFP construct had a subcellular  	
    35	
    pattern very similar to the ER-specific marker p35S:HDEL-RFP (Figure 15B and C), indicating a weak association with the ER both around the nucleus and elsewhere in the cytosol.  3.2.7	
  Stable	
  transformation	
  of	
  p35S:SCD2-­‐GFP	
  in	
  Arabidopsis	
  scd2-­‐1	
  mutants	
  	
   In the last experiment, I wanted to confirm the previous result of ER localization of SCD2 by stable expression in Arabidopsis. To do this, I transformed the p35S:SCD2-GFP constructs into homozygous scd2-1 Arabidopsis mutants and analyzed 1-week old seedlings at the T2 generation. The p35S:SCD2-GFP construct restored the mutant phenotype in Arabidopsis scd2-1 leaves (see Section 3.2.2).  I examined the fluorescence of GFP by laser scanning confocal microscopy. Clear images of localization were produced by observing root cells rather than leaves that have autofluorescing chloroplasts (data not shown). Strong GFP fluorescence was observed in the ER. The reticulate pattern characteristic of the ER was further ascertained by Z stacks of transgenic roots (Figure 16A). Additionally, the cells of the root tip (Figure 16B) and root hairs (Fig 16C) also displayed a strong GFP signal in a characteristic reticulate pattern. One particular epidermal cell was chosen to study the reticulate pattern more closely, and it showed a reticulate pattern (Figure 16D-G). Finally, to confirm the proposed ER-specific localization, roots of transgenic seedlings were stained with hexyl rhodamine B, and SCD2 co-localization with this ER-specific marker was observed (Figure 16H-J).  3.3	
  Discussion	
   Between the previously published patent (Zeng and He, 2010) and two publications reporting on At4g37470 (SCD2) it was not clear (due to data quality) whether SCD2 has a role in wax biosynthesis. To obtain reliable data, I have analyzed cuticular wax of leaves of two independent 	
    36	
    mutant lines scd2-1 and scd2-2, and performed a full characterization of SCD2. All the data acquired in the current work, apart from promoter-GUS analysis, supported the idea that SCD2 is a component of the alkane-forming machinery required for modifying aldehydes to alkanes. This adds yet another gene challenging the previously reported simple two-step pathway hypothesis of alkane production in Arabidopsis (Cheesbrough and Kolattukudy, 1984).  In order to implicate SCD2 in alkane-forming branch of wax biosynthesis, its mutant(s) would be expected to have a decrease in alkanes and an increase in aldehydes (Jenks et al., 1995; Bourdenx et al., 2011; Bernard et al., 2012). Indeed, I observed this in two independent mutant lines of scd2-1 and scd2-2, with the latter having a greater decrease in alkanes compared to its Ler wild type, indicating that the Ds mutation in the scd2-2 allele is a stronger mutant. Neither I nor Zeng and He (2010) observed changes in the stem wax composition compared to the wildtype levels. This indicates that SCD2 is involved in converting aldehydes to alkanes only in Arabidopsis leaves.  To further confirm the involvement of the SCD2 gene in wax biosynthesis, the cDNA of SCD2 was driven under the control of the native SCD2 promoter and expressed in the scd2-1 mutant background. The scd2-1 mutant was chosen for transformation, as its wild-type background Col7 produces many transformants, as opposed to the Ler background of scd2-2 (Cough and Bent, 1998). Two complementation lines were analyzed and only one representative line (pSCD2:SCD2 in scd2-1) is shown. The complementation line indeed restored the alkane and aldehyde levels to those seen in the corresponding Col-7 wild type. This result confirmed the involvement of SCD2 in alkane biosynthesis. In addition, the p35S:SCD2-GFP line also complemented the scd2-1 leaf wax phenotype to that of Col-7. Both of these complementation lines provide evidence that the decrease in alkanes and increase in aldehydes of scd2-1 rosette 	
    37	
    leaves was due to the mutation in the SCD2 gene. My analyses of the two mutant lines and the transgenics clearly show SCD2’s participation in wax biosynthesis.  The cutin monomer composition of scd2-1 did not differ (Zeng and He, 2010) from wild-type levels in leaves and stems, ruling out any participation of SCD2 in cutin biosynthesis. In contrast, another putative wax biosynthetic gene involved in alkane formation, the long chain acyl-CoA synthetase (LACS) CER8, had a mutant phenotype with a characteristic decrease in alkanes, as well as altered cutin composition in Arabidopsis (Lu et al., 2009).  It is very interesting that the strong wax chemical phenotype of scd2 is highly organ-specific and occurs in rosette leaves as opposed to inflorescence stems, which is atypical for mutants implicated in wax biosynthesis. The SCD2 transcript was detected quantitatively in several organs, a finding that is in line with the publically available Bio-Array resource eFP Browser data (University of Toronto). The highest transcript abundance was seen in the rosette and less in cauline leaves. This data is in agreement with the observed wax chemical data in rosette leaves. In contrast, stems had an approximately 50% reduction in SCD2 transcript abundance than did leaves. This indicates that SCD2 is very weakly expressed in stems, matching the results that mutant stem wax exhibited no discernable change from wild type in any of the wax chemical classes. Interestingly, SCD2 transcript was at a considerable level detected in siliques, indicating a possible role for SCD2 in silique cuticle biosynthesis as well. Additional work is required to determine the silique cuticular wax profile to better understand the possible role of SCD2 in silique cuticle biosynthesis. The lowest SCD2 transcript abundance was in roots of four-week old Arabidopsis plants, suggesting that SCD2 is not likely involved at this mature stage in synthesizing a protective lipid layer. This is in contrast to multiple cuticle-related genes whose transcripts were detected in the root tissue, such as FDH (Yephremov et al., 1999; Pruitt et al., 	
    38	
    2000), LCR (Wellesen et al., 2001), WAX2/YRE/CER3 (Kurata et al., 2003), LACS2 (Schnurr et al., 2004), ACE/HTH (Krolikowski et al., 2003; Kurdyukov et al., 2006a), CER5 (Pighin and Zheng et al., 2004) and BDG (Kurdyukov et al., 2006b). However, the GUS-promoter data showed that, at an early developmental stage (two-week old seedlings), SCD2 is expressed in roots, suggesting that in the developing seedling SCD2 is involved but is down-regulated during the development of the seedling towards maturity.  Sun et al (2011) analyzed transgenic plants harboring the pHTL:GUS construct and found that the promoter activity was in the vasculature. However, no explanation or hypothesis as to what the function of HTL may be was provided. The results in this chapter confirm the vasculaturespecific expression of SCD2. My examination of the promoter activity of SCD2 further revealed that it is absent from the xylem, but promoter activity was detected adjacent to it, in the phloem of the vasculature. This finding came as a surprise, as all previously characterized wax-related genes have promoter activities in epidermal cells, leaving SCD2’s precise role in the vasculature without a clear explanation. For extra confirmation, a GUS-promoter fusion of 1.5 kb or 2 kb could be made in the future to examine if the previously seen vasculature expression is also found for the longer version of the promoter, or if it has an epidermal cell expression more consistent with the location of wax biosynthesis. Until then, a clear function cannot be assigned. Previously characterized wax biosynthetic proteins have been localized to the ER in planta, Specifically, MAH1 (Greer et al., 2007) and the fatty elongase components CER6 and CER10, (Kunst and Samuels, 2003; Zheng et al., 2005) were localized to the ER in planta. Apart from these proteins, only CER4 was localized to the ER in yeast, but never confirmed in planta. In the current work, I demonstrate that SCD2, a predicted thioesterase/hydrolase, is localized to the ER in transgenic Arabidopsis plants harboring the p35S:SCD2-GFP construct, which as previously discussed is the location of the fatty acid elongation and subsequent functionalization of wax 	
    39	
    constituents. This provides yet another line of evidence for SCD2’s involvement in the wax biosynthetic pathway.  Although SCD2’s amino acid sequence is homologous to proteins of the alpha/beta hydrolase/thioesterase family, its precise function as a possible enzyme in alkane formation is unknown. The current work has provided a good foundation to continue to characterize SCD2 biochemically. However, SCD2’s localization to the ER can already give some clues to its function. The following two possibilities seem feasible:  One possibility is that SCD2 is associated with the ER membrane through post-translational attachment of a lipid on the cytosolic side of the ER, anchoring SCD2 to it. This scenario describes SCD2 as not participating in an enzyme complex, but functioning as a single enzyme. This might be downstream of the LACS1 (CER8) that first adds a CoA group onto fatty acyl chains as they leave the plastid, and SCD2, a hydrolase/thioesterase localized to the ER removes the CoA group allowing the fatty acid to be further functionalized. The evidence for this comes from SCD2 co-localizing to the ER in Arabidopsis transgenics harboring the p35S:SCD2-GFP construct.  The second possibility is that SCD2 is associated with the ER membrane via protein-protein interactions, possibly with CER3 and/or CER1. The hydrophobicity results show a lack of any trans-membrane domains in the amino acid sequence of SCD2 (Figure 3.10). First evidence for a possible protein complex comes from the subcellular localization of transiently expressing N. benthamiana cells (Figure 15). In that experiment, SCD2 was found weakly associated with the ER on its cytosolic side.  	
    40	
    SCD2 may function as a decarbonylase. Indeed, the phenotypes of scd2-1 and scd2-2 point to the protein having a similar function as CER1, performing the aldehyde decarbonylation by catalyzing the conversion of aldehydes to alkanes. This implies that SCD2 may have the same enzymatic function as CER1. However, there is also strong evidence against a function as a decarbonylase, as SCD2 does not share any homology with the amino acid sequence of CER1 or any of the wax biosynthetic genes. Furthermore, the decarbonylase function has also not been confirmed for CER1 (Bernard et al., 2012; Bernard and Joubes, 2013). Thus, it is difficult to conclude SCD2’s function as a decarbonylase. SCD2’s amino acid sequence does however show similarity with an alpha/beta hydrolase domain, which provides a domain level prediction to SCD2’s function as a hydrolase.  Clearly, both scenarios assign a function to SCD2 and demonstrate a need to further investige SCD2’s enzymatic activity experimentally. This will involve purifying the protein from transgenic E. coli and performing assays testing possible substrates to reveal SCD2’s role as an enzyme. First, the solubility and characteristics of SCD2 will have to be investigated in E. coli prior to subsequent use of the protein in biochemical assays. There are at least two possibilities for enzymatic reactions that can be tested: one is that SCD2 catalyzes a possible hydrolysis of a CoA group to form a fatty acid. Second, SCD2 may perform a decarbonylation reaction removing a CO from the aldehyde and producing the alkane product. Fundamentally, these two reactions are different. Finally, SCD2 could be performing a completely different reaction.  Until the enzymatic studies are completed, the characteristics and function of SCD2 can only be hypothesized and speculated based on the leaf wax phenotype and the properties of the amino acid sequence. Nevertheless, the localization results described earlier in this chapter provide more validation for SCD2 having a role as a hydrolase and not a decarbonylase since CER1 has 	
    41	
    trans-membrane domain topology to the ER and SCD2 does not, thereby providing a line of evidence for SCD2’s possible participating in an enzyme complex. In addition, the fact that the SCD2 protein sequence lacks signaling domains points to its localization to the ER being mediated by protein-protein interactions to function as an enzyme in a protein complex. Thus, combined with the strong chemical phenotype in leaves with the localization results, there is more evidence than before supporting SCD2’s involvement as an additional component of the alkane-forming machinery in Arabidopsis leaves.  	
    42	
    Figure 5 Cuticular wax composition of leaves and stems of Arabidopsis Col-7 (WT) and scd2-1. Wax coverage is expressed as (µg/cm2). Mean values are given with SD (n=3). Each wax constituent is designated by carbon chain length and is labeled by chemical class along the xaxis. Taken from Zeng and He (2010). Abbreviations Alk=alkane, PA=primary alcohol, ALD=aldehyde, FFA=free fatty acid, KE=ketone. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    43	
    	
   Figure 6 Cutin monomer composition of leaves and stems of Arabidopsis Col-7 (WT) and scd21. Monomer amounts are expressed as (µg/cm2) of stem and leaf surface. Mean values are given with SD (n=3). Adapted from Zeng and He (2010).  	
    44	
    Col -­‐7  40  scd2-­‐1  %	
  of	
  t otal	
  stem	
  wax  35  Ler  30  scd2-­‐2  25  pSCD2::SCD2	
  in	
  scd2-­‐1 p35S::SCD2-­‐GFP	
  in	
  scd2-­‐1  20 15 10 5  Not	
  identified  Triterpenoids  Compound	
  classes  Esters  Primary alcohols  Ketones  Secondary alcohols  Alkanes  Aldehydes  Fatty	
  acids  0  	
   	
   Figure 7 Cuticular wax composition on inflorescence stems of Col-7 (WT), scd2-1, Ler, scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major compound classes are expressed as (% of total stem wax) of stem surface area. The data represent mean values ± SD of n=5. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    45	
    120 Col-­‐7 scd2-­‐1  100  Ler scd2-­‐2  30  scd2-­‐2  scd2-­‐2  25  pSCD2::SCD2	
  in	
  scd2-­‐1  pSCD2::SCD2-­‐GFP	
  in	
  scd2-­‐1 p35S::SCD2-­‐GFP	
  in	
  scd2-­‐1  pSCD2::SCD2	
  in	
  scd2-­‐1 p35S::SCD2-­‐GFP	
  in	
  scd2-­‐1  p35S::SCD2-­‐GFP	
  in	
  scd2-­‐1  20 15 10 5  Not	
  identified  Triterpenoids  Esters  Primary alcohols  0 Ketones  00  Ler  Secondary alcohols  20  Ler  Alkanes  20  scd2-­‐1  Aldehydes  40  scd2-­‐1  Fatty	
  acids  60  40  Col-­‐7  35  100  6080  Col -­‐7  40  80  %	
  of	
  t otal	
  stem	
  wax  %	
  of	
  compound	
  class %	
  of	
  compound	
   class  120  2828 3030 2828 30 27 29 31 31 33 33 24 2626 2828 3030 3232 30 27 29 24 40 40 42 42 44 44 46 46 48 4850 50 Fatt	
  aacid	
   Aldehyde	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  Alkane	
   Primary	
   alcohol	
   Alkane	
   Fatty	
   cid	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  Aldehyde	
   	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  Primary	
   alcohol	
   	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  EEster	
   ster Compound	
  classes Chain	
  lChain	
   ength	
   of	
  compound	
   class length	
   of	
  compound	
   class  	
   Figure 8 Cuticular wax composition on inflorescence stems of Col-7 (WT), scd2-1, Ler, scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major components are expressed as (% of compound class) of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=5. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    46	
    140  120  Wax	
  coverage	
  [µg/	
  dm 2 ]  100  80  60  40  20  0 Col-­‐7  scd2-­‐1  Ler  scd2-­‐2  pSCD2::SCD2	
  i n scd2-­‐1  p35S::SCD2-­‐GFP	
  i n scd2-­‐1  	
   Figure 9 Cuticular wax coverage on rosette leaves of Col-7 (WT), scd2-1, Ler (WT), scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Total wax coverage is expressed as µg/dm2 of leaf surface area. The data represent mean values ± SD of n=5. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    47	
    Table 1 Cuticular wax composition of rosette leaves of Arabidopsis Col-7 (WT), scd2-1, Ler (WT), scd2-2, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3).  Fatty acids Aldehydes Alkanes Primary alcohols Not identified Total  Col-7 3.6 ± 0.18  scd2-1 3.4±0.16  Ler 3.5±0.20  scd2-2 3.6±0.22  pSCD2:SCD2 in scd2-1 3.2±0.13  p35S:SCD2GFP in scd2-1 3.3±0.19  13.3 ±0.79 55.4 ±3.87 26.7 ± 1.87 23.1 ± 1.15 119.5 ±11  30.6±2.14 35.9±2.83 24.1±1.44  12.4±0.99 47.1±4.23 23.8±1.61  28.1±2.24 14.5±1.16 24.6±2.09  14.1±1.01 51.0±4.02 24.1±1.80  13.8±1.24 53.2±3.71 25.9±1.68  18.0±1.53  23.2±1.74  21.5±1.39  27.7±1.52  22.9±1.71  112.0 ±8.0  110 ± 10  92.3 ± 9.5  120.0 ± 12.0  119.0 ± 10.5  	
    	
    48	
    35  Wax	
  coverage	
  [µg/	
  dm 2	
  ]	
    30 25  Col-­‐7 scd2-­‐1  20  Ler scd2-­‐2  15 10 5  br-­‐34  br-­‐32  n-­‐34  br-­‐30  n-­‐32  n-­‐30  n-­‐28  Alkane  Chain	
  length	
  by	
  compound	
  class  37  35  33  31  29  27  34  32  30  26  24  34  28  Aldehyde  n-­‐26  Fatty	
  acid  32  30  28  26  24  0  Primary	
  alcohol  Figure 10 Cuticular wax composition on rosette leaves of Col-7 (WT), scd2-1, Ler (WT), and scd2-2 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the xaxis. The data represent mean values ± SD of n=5. br=branched primary alcohols.  	
    49	
    35 Col-­‐7  30  Wax	
  coverage	
  [µg/	
  dm 2	
  ]  scd2-­‐1 pSCD2::SCD2	
  in	
  scd2-­‐1  25  p35S::SCD2-­‐GFP	
  in	
  scd2-­‐1  20 15 10 5  br-­‐34  br-­‐32  n-­‐34  br-­‐30  n-­‐32  n-­‐30  n-­‐28  Alkane  Chain	
  length	
  of	
  compound	
  class  37  35  33  31  29  27  34  32  30  26  24  34  28  Aldehyde  n-­‐26  Fatty	
  acid  32  30  28  26  24  0 Primary	
  alcohol  Figure 11 Cuticular wax composition on rosette leaves of Col-7 (WT), scd2-1, pSCD2:SCD2 in scd2-1 and p35S:SCD2-GFP in scd2-1 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=5. br=branched primary alcohols.  	
    50	
    Figure 12 Differential expression analysis of Arabidopsis SCD2 in various organs of four-weekold Arabidopsis plants. The gene expression level was determined by real-time RT-PCR analysis. Results are represented as relative transcript abundances. The data represent the means ± SD of three biological and two technical replicates (n=6). Total RNA was isolated from roots, rosette and cauline leaves, bottom- mid- and top section of stem, flowers and siliques. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    51	
    	
   	
   Figure 13 Spatial expression pattern the SCD2 gene in transgenic Arabidopsis plants harboring the SCD2 promoter fused to the GUS gene. Promoter activity was visualized through histochemical GUS staining on two-week old T2 transgenic seedlings. A and B. Whole seedling imaged by dissecting light microscope, C and D. mature leaves, E. hypocotyl F. petiole and shoot apex, G. emerging true leaf, H. root tip and differentiation zone, I. secondary roots, J and K. DIC image of root, L. True leaf mid-vein. A-B bars= 5mm, J and L bars=20 µm, K bar=15 µm ,C-I bars=0.2 mm. 	
    52	
    	
   	
   Figure 14 Hydrophobicity plots of various wax biosynthetic proteins. A. SCD2 B. CER8 C. CER2 D. CER6. Data generated using amino acid sequences of various proteins in the TMHMM program. 	
   	
   	
    53	
    A	
    	
    	
    	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  	
  B	
    	
    	
    	
  	
  	
  	
  	
  	
  	
  	
  C	
    	
   	
   Figure 15 Transient expression of p35S:SCD2-GFP and p35S:HDEL-GFP in N. benthamiana epidermal cells. A. p35S:SCD2-GFP, B. p35S:HDEL-GFP, and C. as merge. Bars=2µm. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    54	
    	
    	
   Figure 16 Subcellular localization of SCD2 in root epidermal cells of stably transformed Arabidopsis scd2-1. Confocal microscopy images of p35S:SCD2-GFP. A. Z stack of root. B. root tip. C. root hairs and root epidermal cells. D, E, F, G. Planes through the root epidermal cell. H. root epidermal cell under 488 nm excitation (GFP). I. hexyl rhodamine stain under 561 nm excitation (RFP). J. merge. Bars=20µm A-G.  	
    55	
    Chapter	
  4	
  Characterization	
  of	
  cer1cer3	
  and	
  elucidation	
  of	
  epistatic	
   relationships	
  of	
  CER	
  genes	
  in	
  Arabidopsis	
  wax	
  biosynthesis	
   	
    4.1	
  Introduction	
   Progress in understanding the involvement of various genes in the wax biosynthetic pathway has been hampered by the inability to accurately predicting gene product functions based on the sequence homology to known proteins. Many CER genes have been identified based on the mutant chemical phenotypes. To advance our knowledge on the gene interactions that control wax biosynthesis, various double mutants have been studied previously (Goodwin et al., 2005). In that study, 11 homozygous recessive cer mutants were used to create 14 double cer mutants. The Wassilewskija (WS) cer1-cer4 alleles used in Goodwin et al (2005) were produced using TDNA insertion mutagenesis as described in McNevin et al (1993). In that study, the cer1cer3, cer1cer4 and cer3cer4 double mutants had a drastic reduction of total stem wax load, but not lower than either of the respective parents. The results of the three double mutant lines can be summarized as follows: when cer1 was placed in the cer3 background (mutant with primary alcohol amounts much higher than in the wild type), the primary alcohol levels were much lower than in cer3, specifically the C30 primary alcohol was intermediate to the parents. The cer4 mutation blocks the conversion of acyl-CoAs to primary alcohols. The cer1cer4 mutant produced significantly more primary alcohols and their metabolites (esters) than the cer4 mutant. Lastly, when cer3 was placed in the cer4 background, the cer3cer4 double mutant resembled the cer4 single mutant by having extremely low levels of primary alcohols. In all cases, levels of alkanes were dramatically decreased in the double mutants, but cer1cer4 had the least amount compared to both parental lines.  	
    56	
    Although the double mutant study by Goodwin et al (2005) presented interesting results, there are weaknesses that cannot be overlooked. First, none of the lines were genotyped in Goodwin et al (2005), but rather were selected based on a glossy stem phenotype. To find double mutants, these were examined using gas chromatography (GC) to select putative doubles having wax composition different from both cer mutants. Since each mutation has a recessive mode of inheritance, there is a risk of incorrectly identifying double homozygous mutants based on a glossy stem phenotype. One cer mutation can be homozygous at one locus, while the second cer gene can be either heterozygous or homozygous for the mutation. Both of these possible genotypes will have a glossy stem phenotype. Therefore, it is important to PCR-genotype all glossy plants to properly identify the true double mutants for use in subsequent cuticular wax chemical analysis. Second, the cer1 parent mutant did not have elevated C30 aldehyde levels, which differed from previous results of the same group (Jenks et al., 1995), and therefore puts the quality of the data in question. Third, cer mutants have been available only in the WS or Ler backgrounds having T-DNA insertions or EMS-based mutations and many were not null. Fourth, this study of epistatic relationships was only performed on inflorescent stems while double mutant leaf characterization is lacking. An investigation of leaf wax of double mutants is important for furthering the understanding of leaf wax biosynthetic pathway.  Therefore, for the reasons described above, I have selected available published mutants in the Col-0 background that are null cer1-2 (Bourdenx et al., 2011) and cer3-6 (Kurata et al., 2003; Rowland et al., 2007), and cer4-3 (Rowland et al., 2006) based on the published RT-PCR data. From here on, cer1-2, cer3-6 and cer4-3 will be denoted as cer1, cer3 and cer4, respectively. I generated three double mutant combinations cer1cer3, cer1cer4 and cer3cer4, all of which were PCR-genotyped and analyzed for perturbations in both stem and leaf cuticular waxes. These three particular combinations were chosen to study the branch-point of the wax biosynthetic 	
    57	
    pathway, where the elongated acyl-CoAs are channeled into the primary alcohol pathway or the alkane-forming pathway. In particular, the following questions were addressed: Is there clear epistasis in any of the double mutant lines? If so, which genes are sequential and which belong on separate branches of the pathway?  Further motivation for producing the cer1cer3 double mutant was: first, to generate a double mutant tool with stronger alleles than were crossed and reported previously by Goodwin et al (2005) to test in planta the various constructs of truncations and fusions of CER1/CER3 that I generated (See Chapter 1 for a review on CER1 and CER3 domain structure); and second, to characterize the double mutant in more detail including wax crystal coverage on stems and to investigate both qualitatively and quantitatively the male-sterility phenotype.  4.2	
  Results	
   All single mutants cer1, cer3 and cer4 had a bright green, glossy stem phenotype. The cer1cer3, cer1cer4 and cer3cer4 also had a glossy stem. The characterization of the cer1cer3 was carried out in detail and is described first, followed by chemical analyses of the three double mutant combinations.  4.2.1	
  Generating	
  the	
  cer1cer3	
  double	
  mutant	
   The cer1cer3 double mutant was generated by crossing cer1 and cer3. Since the cer3 single mutant was semi-sterile, I tried two different combinations of donor and recipient (Figure 17) for producing viable progeny. A total of three viable F1 seeds were harvested. Each homozygous double mutant gave a total of 9 seeds.  	
    58	
    4.2.2	
  The	
  cer1cer3	
  double	
  mutant	
  exhibits	
  a	
  glossy	
  stem	
  phenotype	
   One of the characteristics of a wax-deficient Arabidopsis mutant is a glossy stem phenotype that indicates a reduction in wax coverage. Both cer1 and cer3 parental lines had a glossy, bright green stem compared to the wild type, which has a glaucous or whitish appearance and pale green color indicating normal levels of wax (Figure 18A). The cer1cer3 double mutant had a bright green and glossy stem phenotype.  4.2.3	
  The	
  cer1cer3	
  double	
  mutant	
  exhibits	
  a	
  severe	
  additive	
  sterility	
  defect	
   The wild-type siliques are very long and straight, as well as having a glaucous appearance. The cer3 single mutant has a curled silique phenotype as well as a great reduction in seed set. The cer1 siliques are straight, glossy and long, cer3 siliques are shorter, glossy and curly. To study whether the double mutant has the same affliction, a representative photograph was selected to show the phenotype (Figure 18B). The cer1cer3 double mutant has glossy, shorter, and curlier siliques. The cer1 plants were more fertile than cer3 under increased conditions of humidity, while cer1cer3 was severely sterile under the same conditions.  To quantitatively measure the sterility phenotype, a seed count per plant was performed on wildtype, cer1, cer3 and the cer1cer3 double mutant (Figure 18C). Wild-type plants produced 5,000 (± 50) seeds per plant, cer1 4,500 (± 50) seeds, cer3 only 120 seeds (±10), and the cer1cer3 double mutant only 9 seeds, indicating a severe sterility phenotype.  4.2.4	
  The	
  cer1cer3	
  double	
  mutant	
  has	
  a	
  reduction	
  of	
  wax	
  crystals	
   To provide a quantitative analysis of the glossy stem phenotype of the cer1cer3 double mutants, scanning electron micrographs (SEM) were taken of the top 1 cm stem segment of wild type, single and double mutants to compare epicuticular wax crystal coverage (Figure 19). The 	
    59	
    epicuticular wax coverage was high in wild type. The single cer1 mutant was devoid of wax crystals. The cer3 single mutant had a substantial reduction in wax crystals compared to wild type. The cer1cer3 double mutant had similar epicuticular crystal presence, as per the cer3 mutant. 	
   4.2.5	
  Arabidopsis	
  cer1,	
  cer3	
  and	
  cer1cer3	
  show	
  altered	
  cuticular	
  wax	
  composition	
   To investigate the effect of CER1 and CER3 mutations on cuticle composition, the wax of fourweek-old Arabidopsis stems and leaves of Col-0 (wild type, WT), cer1, cer3 and cer1cer3 was analyzed in detail (Figures 20 and 23; Tables 2 and 3).  In stems, the wax loads of cer1 (255.0 ± 15.3 µg/dm2) and cer1cer3 (319.2 ± 22.3 µg/dm2) were drastically reduced by approximately 8-fold, and in cer3 (781.3 ± 46.8 µg/dm2) by approximately 2-fold compared with wild-type (1622.0 ± 97.3 µg/dm2) plants (Table 4.1). The decrease measured in cer1cer3 was largely due to reduced levels of the three major stem wax compounds: C29 alkane, C29 secondary alcohol and C29 ketone (Table 2 and Figure 20). Compared to cer3, cer1cer3 had a similar chain length distribution of primary alcohols. The C30 primary alcohol was increased by 4-fold in cer1cer3 and 5-fold in cer3 compared to wild type. The level of aldehydes was greatly reduced in cer1cer3 and resembled those measured in cer3, rather than cer1 (Figure 20). Interestingly, the C24 fatty acid was greatly increased in cer1cer3 while the levels in wild type and parental mutants were barely detected (Figure 20). The C26 fatty acid was drastically reduced in both single and double mutants compared with wild type. Finally, the quantity of esters measured in cer1cer3 was increased by 38%, while the single mutants were unchanged from wild type (Table 2). The rise in esters is likely due to the presence of the higher levels of C28 and C30 primary alcohol in the double mutant, as they are precursors for esters. 	
    60	
    In leaves, the wax load was also reduced (as in stems) albeit to a milder extent, with a two-fold decrease from wild-type (142.2 ± 11.3 µg/dm2) to cer1 (71.7 ± 6.4 µg/dm2), cer3 (79.8 ± 5.5 µg/dm2), and cer1cer3 (67.1 ± 5.3 µg/dm2) (Table 3). The greatest reduction was due to alkanes (Figure 23). The cer1cer3 had the lowest amount of C29 alkane and had an intermediate level of C31 and C33 alkanes compared to both single mutant parents. The cer1cer3 alcohol (C26-C32) and aldehyde (C30-C34) amounts resembled cer3 rather than cer1. No major differences were seen in the branched alcohols. The cer1cer3 double mutant had an increase in the C24 fatty acid level, similar to stems. The C26 fatty acid was decreased in all mutants compared to the wild type. The C30 fatty acid showed a 2.5-fold increase in cer1 compared to wild-type, while in cer3 and cer1cer3 it was not detected.  4.2.6	
  Arabidopsis	
  cer1cer4	
  and	
  cer3cer4	
  show	
  altered	
  cuticular	
  wax	
  composition	
   The wax load of cer1cer4 stems (178.9 ± 12.52 µg/dm2) was greatly reduced relative to wild type, even beyond the cer1 (255.0 ± 15.30 µg/dm2) parent (Table 4.1). This great reduction was due to the lack of C29 alkane, C29 secondary alcohol and C29 ketone (Table 2 and Figure 21). The levels of both primary alcohols and esters were intermediate between cer1 and cer4. Interestingly, the primary alcohols of the double mutant were dominated by chain length C28. The C26 fatty acid was reduced in both single mutants, but was found at equivalent levels in the double and the wild type (Figure 21). Interestingly, C30 aldehyde was elevated in cer1 and more so in cer4, while it was equivalent in cer1cer4 to wild type.  In leaves, the wax load was also reduced in cer1cer4 (57.4 ± 3.9 µg/dm2) beyond that of cer1 (71.7 ± 6.4 µg/dm2) (Table 3). The greatest reduction in wax load came from a loss of C29, C31 and C33 alkanes in cer1cer4 (Figure 24). The alkane levels of those chain lengths in the double 	
    61	
    mutant were similar to cer1. The C28 aldehyde was present in wild type, cer1 and cer4, but was not detected in cer1cer4. The C30, C32 and C34 aldehydes were slightly elevated in cer1 and cer4, but were unchanged in cer1cer4 compared to wild type. The C24 fatty acid was elevated beyond the wild type and single mutants. The C26 fatty acid was reduced in the single mutants, but was equivalent in cer1cer4 to wild type. The C30 and C32 fatty acids were elevated in cer4 and more so in cer1, but were reduced in the double beyond both single mutant parents. The C28 primary alcohol levels in the double resembled the cer1 mutant, whereas C26 and C30 were not present. No significant changes were detected in branched alcohols.  In stems, the wax load of cer3 (781.3 ± 46.8 µg/dm2) was greatly reduced compared to wild type (1622.0 ± 97.3 µg/dm2), while cer4 (1591.2 ± 111.3 µg/dm2) was unchanged. The cer3cer4 (275.1 ± 22.1 µg/dm2) double mutant had greatly reduced total wax load, beyond cer3 (Table 2). The decrease in wax load was due to a loss of C29 alkane, C29 secondary alcohol and C29 ketone (Figure 22 and Table 2). Interestingly, the C31 alkane level in cer3 and cer3cer4 was elevated 2-fold compared to wild type (Figure 22). The aldehydes and fatty acids were the least abundant of all compound classes in cer3cer4, similar to cer3. The amounts of primary alcohols and esters were extremely reduced in the double mutant. The double mutant had extremely reduced amounts of C30 primary alcohols, resembling the cer4 parent mutant.  In leaves, the wax load of the cer3cer4 double mutant (56.3 ± 5.0 µg/dm2) was reduced compared to cer3 (79.8 ± 5.5 µg/dm2), even though it was unchanged between cer4 (120.1 ± 10.8 µg/dm2) and wild type (Table 3). The levels of fatty acids were extremely low in cer3cer4, similar to the respective single mutants (Figure 25). The levels of aldehydes in cer3cer4 were similar to cer3 (C30-C34). The double mutant levels of alkanes (C29-C33) were similar to cer3. 	
    62	
    The primary alcohols (C26-C30) were dramatically reduced in the double mutant, similar to the cer4 parent. Levels of branched primary alcohols were unchanged.  4.3	
  Discussion	
   Although wax biosynthesis has been extensively studied in the model plant Arabidopsis, the precise functions of many gene products are not yet confirmed. Evidence for their function comes from characterizing Arabidopsis single mutant wax chemical profiles that show a simple increase in a particular chemical constituent and a subsequent decrease in a downstream one, thereby pointing to a likely enzymatic reaction that synthesizes a particular product from a substrate. However, this relatively simple chemical phenotype from a single cer mutant becomes more complicated when coupled with a second cer mutant. As previously introduced (section 4.1), Goodwin et al (2005) generated 14 cer double mutant combinations whose chemical phenotypes in Arabidopsis inflorescent stems varied greatly and displayed the same, intermediate, or lower/higher levels of individual compounds compared to the respective single mutants. However, many weaknesses were present in that study that put the quality and reliability of the results in question. To study wax biosynthesis more accurately, I used stronger alleles where available, generated my own double mutant lines and PCR-genotyped them (Figure 17C) and re-analyzed the chemical composition of both stems and leaves. Most importantly, the characterization in leaves as presented in this thesis is the first and only report of its kind.  This discussion first focuses on the physiological characterization of the cer1cer3 double mutant, followed by stem and leaf wax composition of the three double mutant lines.  	
    63	
    4.3.1	
  The	
  cer1cer3	
  double	
  mutant	
  is	
  severely	
  male-­‐sterile	
   Upon visual examination of the cer1cer3 mature plants of four-weeks of age, few siliques were present. The severe sterility of cer1cer3 was further characterized quantitatively by counting seed set per plant. A single cer1cer3 plant produced only nine seeds and had severely curly and short siliques. The cer1 plants are more fertile than cer3 under increased humidity conditions, while cer1cer3 was severely sterile under the same conditions. Thus, mutations in both genes caused a more severe fertility defect. There are three possible explanations for the fertility defect in the single and double mutant lines.  One possible explanation is that the mutations in CER1 and CER3 affect pollen exine production. A primary component of the exine is sporopollenin, a polymer consisting of aliphatic and aromatic molecules cross-linked by ester and ether bonds making it highly resistant to degradation (Piffanelli et al., 1998; Scott et al., 2004; Ma, 2005). Amongst other roles, the exine facilitates pollen-stigma interactions. The CER3/YORE-YORE/WAX2/FACELESSPOLLEN1 was previously shown to be associated with exine production and male fertility (Ariizumi et al., 2003; Chen et al., 2003; Kurata et al., 2003). SEM revealed that the pollen surface was almost smooth and lacked a reticulate pattern on flp1, hence the name faceless pollen-1 (flp1). SEM also showed that the exine was easily damaged by acetolysis, indicating that sporopollenin was defective in the mutant (Ariizumi et al., 2003). Further characterization of the sterility defect in the cer1cer3 double mutant by SEM could shed more light onto the physical structure of the pollen grains. It is likely that its severe male sterility is due to a compromised exine that in turn hindered pollenstigma interaction preventing successful fertilization (Zinkl et al., 1999; Zinkl and Preuss, 2000).  Another possible explanation for the male sterility of cer3 and cer1cer3 is an abnormal exine structure and a missing hydration signaling component (Preuss et al., 1993) in tryphine. The 	
    64	
    tryphine droplets, which are composed of VLC cuticular compounds, were found to be reduced in cer6, and reduced in size but more numerous in cer1 and cer3 (Preuss et al., 1993; Hülskamp et al., 1995; Aarts et al., 1995). It was shown that cer1, cer3 and cer6 mutants are male-sterile in low humidity conditions but are normally fertile in high humidity conditions (Preuss et al., 1993; Hülskamp et al., 1995; Aarts et al., 1995; Fiebig et al., 2000). In contrast, the cer1cer3 plants produced only nine seeds per plant when grown in high humidity. This result suggests that the probable cause of male sterility may lie in both an abnormal exine structure contributed by the cer3 parent and an altered tryphine in cer1 that lacks a signaling molecule for hydration of the pollen grains.  The third explanation is a compromised cutin in anthers. There may be a cutin defect in cer3, suggested by the curliness of the siliques and also an organ fusion phenotype in the inflorescences and true leaves, although not 100% penetrant. However, a previous study has carried out a detailed investigation of the cutin profile in two different cer3 lines (cer3-6 and cer3-4) and no obvious reduction in any cutin monomer in the leaves was found (Rowland et al., 2007). The conclusion drawn by the authors was that CER3 activity, at least in leaves, is not required for cutin biosynthesis. However, this may not be true for other tissues and organs since the WDA1 protein from rice that is related to CER1 and CER3 was previously reported to be involved in cutin monomer formation in the anthers of rice. This may suggest that CER3 might have an analogous role in anthers and pollen grains. Since the mutations have an additive sterility phenotype, this may indicate that the roles of CER1 and CER3 are somewhat redundant in anther and pollen grain cutin biosynthesis.  Finally, the male sterility of cer1cer3 may be due to all the afore-mentioned factors if not entirely then at least partially. 	
    65	
    4.3.2	
  Wax	
  crystals	
  are	
  present	
  on	
  cer1cer3	
   A full characterization of the double mutant in this study includes documenting the glossy stem phenotype not previously reported. The glossiness of the double mutant was not unexpected, since both the parental lines have a glossy stem. This indicates that the double mutant also has a great reduction in wax coverage on the surface of the stem. This was confirmed by subsequent analysis of the stems by SEM. The cer1cer3 double mutant had a reduction in wax crystals like the cer3 mutant, but not a complete reduction as was observed in cer1. It is known that the formation of epicuticular wax crystals is due to the presence of C29 alkane, C29 secondary alcohols and C29 ketone (Jetter et al., 2006). The cer1cer3 double mutant had the lowest C29 alkane levels compared to cer1 and cer3. The C29 secondary alcohol and C29 ketone levels were similar to cer3 rather than cer1. Crystallization requires the presence of sufficient amounts of a particular constituent. The double mutant had more secondary alcohols and ketones than cer1, and this may have facilitated crystal growth that was visualized by SEM.  4.3.3	
  CER3	
  precedes	
  CER1	
  in	
  both	
  stems	
  and	
  leaves	
   Total stem wax was extracted from 4 week-old wild type, cer1, cer3 and cer1cer3. Wax chain length distribution displayed a pattern typical for cuticular wax mixtures. Compared to cer1, both cer3 and cer1cer3 had the same reduction in C30 aldehyde levels. The predominant constituent present in wild type stems is C29 alkane. The C29 alkane levels are greatly reduced in all the mutants compared to the wild type. Metabolites of alkanes (secondary alcohols and ketones) were also reduced in the single and double mutants. The C30 primary alcohol levels in stems were altered as well. Specifically, cer1 had a slight reduction, while cer3 and cer1cer3 mutants had a significant increase in C30 primary alcohols (440% and 306% compared to wild type, respectively).  	
    66	
    Taking into account the level of aldehydes, alkanes and primary alcohols in stems and leaves, the order of genes is CER3 first and CER1 second on the alkane-forming branch, since cer1cer3 exhibited a rise in the C30 primary alcohol levels, but slightly lower than cer3. If cer1cer3 showed C30 aldehyde levels similar to cer1, a different sequence would be true where CER1 precedes CER3, but that is not the case.  When cer1 was crossed with cer3 (a mutant with much higher primary alcohol amount than in wild type), the C30 primary alcohol levels on the double mutant were slightly lower than in cer3. Thus, CER1 is required, at least in part, for highest primary alcohol production. At present, it seems that CER1 not only has a role in converting aldehydes to alkanes in both stems and leaves, but also acyl-CoA conversion to primary alcohols. The exact mechanism of the reaction involved is unknown, and thus more biochemical investigations are needed.  It is interesting to note that in the Goodwin et al (2005) publication and in this chapter cer1cer3 had the greatest amount (678% increase compared to wild type) of the C24 fatty acid levels compared to the wild type and respective single mutants. In contrast to the C24 levels, the C26C30 chain lengths were not detected in cer1cer3. The trends are the same in leaves. Although Goodwin et al (2005) report this, they do not discuss this data or speculate on an additional function of CER1 and CER3 in fatty acid elongation. It seems likely that both CER1 and CER3 are necessary for elongation of the C24 acyl-CoA. Until biochemical evidence showing that CER1 and/or CER3 are directly involved in elongation of C24 to C26 acyl-CoA chains is presented, this conclusion is premature.  	
    67	
    4.3.4	
  The	
  cer1cer3,	
  cer1cer4	
  and	
  cer3cer4	
  double	
  mutants	
  reveal	
  complex	
  gene	
  interactions	
   Two of the three double mutants, cer1cer3 and cer3cer4, produced more total wax than at least one of their respective single mutant parents. This likely indicates a compensatory mechanism. One double mutant cer1cer4 produced less total wax than both single mutants. This indicates that both genes are needed to produce wild type amounts of wax. The predominant wax class on Arabidopsis stems and leaves is alkanes. Alkanes and alkane metabolites (secondary alcohols and ketones) had the greatest reduction in all single mutants except cer4. In cer1cer4 and cer3cer4, the total amount of alkanes was significantly lower than in both respective single mutant parents. This implies that the functions of the CER gene products overlap additively in alkane biosynthesis (Goodwin et al., 2005).  The nature of the pathway is unlikely to be linear, but likely contains branches. Some cer mutations that inhibit the flux into the alkane-forming branch cause a redirection of substrates into other branches of the pathway, the extent and direction of which are variable and dependent on the particular mutation. For example, the stems of cer1cer3 had an increase in esters, since an increase in the amount of primary alcohols most likely served as precursors for esters. This was similar to the report in Goodwin et al (2005). Furthermore, cer3 and cer4 represent mutations in the genes at the branch point of the pathway, each receiving acyl-CoA chains to further modify into constituents of the primary alcohol pathway or alkane-forming pathway, respectively. The levels of aldehydes and alkanes in the cer3cer4 double mutant stems resembled those found in cer3 rather than cer4. The levels of primary alcohols and esters were greatly reduced in the cer3cer4 double mutant similar to the cer4 parent. Generally, the chemical phenotype of cer3cer4 matched the phenotype of the individual single mutant parents. Since CER3 and CER4 are generally accepted to govern steps in two separate branches, the order of genes is not sequential. Finally, when cer1 and cer4, two mutants with decreased primary alcohol levels were 	
    68	
    crossed, the cer1cer4 double mutant had an intermediate level between the two, in contrast to Goodwin et al (2005) who found much higher levels on the double mutant compared to both single mutants. The results presented in this chapter suggest that the cer1 mutation does not bypass the cer4 mutation. Further studies are needed to study the involvement of CER1 in the primary alcohol pathway and to which degree it interacts with CER4.  The differences in re-distribution of substrates by the gene mutations shed light on the degree of complexity and multiplicity of operations between the interacting CER genes that control the distribution of substrate flux in the wax biosynthetic pathway. Even though stronger alleles have been used in my work to generate the three double mutant combinations, still no wax class was completely eliminated from the wax fraction. This inability to fully block substrate flux into any branch of the wax biosynthetic pathway either by single or double mutations suggests that there is high redundancy in wax biosynthesis operations.  4.3.5	
  Wax	
  biosynthesis	
  in	
  stems	
  and	
  leaves	
   The cer1, cer3 and cer4 mutants have a wax composition change in both stems and leaves. The results presented in this chapter show that the same trends are present in both stems and leaves of cer1cer3, cer1cer4 and cer3cer4. The mutants cer2 (Jenks et al., 1995; Goodwin et al., 2005; Haslam et al., 2012; Pascal et al., 2013) and cer6 (Jenks et al., 1995; Millar et al., 1999) have a stem-specific wax chemical phenotype without having an obvious phenotype in leaves. However, the double mutant study reported here indicates that CER1, CER3 and CER4 perform similar functions in leaf and stem cuticular wax biosynthesis in Arabidopsis.  Out of the three double mutant lines, cer1cer3, was the only one that displayed a chemical phenotype close to epistasis. The other two double mutant lines, cer1cer4 and cer3cer4, did not 	
    69	
    display clear epistasis. The conclusion from these studies of double mutants is that CER3 precedes CER1 in the alkane-forming pathway, while CER4 belongs to the primary alcohol pathway. Further investigations are required to fully understand the functions of CER1 and CER3. The cer1cer3 double mutant will serve as a tool to test functional complementation of each domain using constructs truncating and fusing these domains that have already been made. Finally, the cer1cer3 line may also be used to investigate the contribution of CER1/CER3 to pollen development.  	
    70	
    	
    	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    C  71	
    Figure 17 Strategy used to cross cer1 with cer3 to produce the cer1cer3 double mutant. A. The successful and B. unsuccessful crossing combinations. C. Genotyping of cer1, cer3, cer4 single mutants and cer1cer3, cer1cer4 and cer3cer4 double mutants. Gene specific primers were used to PCR-amplify the WT locus of CER1, CER3 and CER4. Primers are denoted by abbreviations F=forward primer, R=reverse primer, C1=CER1, C3=CER3, C4=CER4. The Lba1 primer was used in combination with the appropriate forward or reverse primers of the WT locus to check presence of the T-DNA in both single and double mutants.  	
    72	
    Figure 18 Phenotype characterization of the cer1cer3 mutant. A. Visual stem phenotype of Col0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant. B. Visual silique phenotype of Col-0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant. C. Quantitative analysis of sterility by seed count (per plant) from Col-0, cer1-2 and cer3-6 single parental mutants, and cer1cer3 double mutant.  	
    73	
    Figure 19 Scanning electron micrographs of wild-type, cer1-2, cer3-6, and cer1cer3 stems. Bars=2µm. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    74	
    Table 2 Cuticular wax composition of inflorescence stems of Arabidopsis Col-0 (WT), cer1, cer3, cer4, cer1cer3, cer1cer4 and cer3cer4. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3). 	
    Triterpenoids Not identified  Col-0 25.1± 2.25 52.3± 4.18 761.2± 68.5 178.5± 14.24 388.0± 31.04 120.7± 11.86 49.2± 4.43 31.4± 2.51 18.7± 1.49  cer1 15.6± 1.24 81.8± 5.72 14.3± 1.01 5.0± 0.35 9.3± 0.65 40.5± 3.24 45.3± 3.15 28.1± 1.96 15.2± 1.06  cer3 7.3± 0.36 10.9± 0.87 230.2± 18.41 43.0± 3.44 172.6± 15.48 238.8± 19.1 44.5± 3.56 25.3± 1.77 8.70± 0.78  Total  1622.0± 97.32  255.0 ± 15.30  781.3± 46.87  Fatty acids Aldehydes Alkanes Secondary alcohols Ketones Primary alcohols Esters  cer4 16.7± 1.17 149.8± 13.48 843.0± 59.01 156.9± 12.56 365.0± 25.55 13.5± 0.94 10.0± 0.92 26.1± 2.41 10.2± 0.71 1591.2 ± 111.37  cer1cer3  cer1cer4  cer3cer4  11.7± 0.82  18.4± 1.47  1.8±0.16  7.4±0.37 17.2 ± 1.37  48.1± 3.36  5.5±0.27  8.9±0.71  121.2± 9.68  17.0±1.19  6.9±0.41  18.4± 1.47  51.3±6.01  0.3±0.01  71.0±4.97  156.6± 12.48  23.5± 1.41  6.7±0.47  68.2± 6.13  27.6± 2.21  9.3±0.74  27.3± 2.75  30.9± 2.70  28.4± 2.24  13.6± 1.08  14.3± 1.00  12.8± 0.89  319.2± 22.34  178.9± 12.52  275.1± 22.13  	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    75	
    Table 3 Cuticular wax composition of rosette leaves of Arabidopsis Col-0 (WT), cer1, cer3, cer4, cer1cer3, cer1cer4 and cer3cer4. Mean values (µg/dm2) of total wax loads and coverage of individual compound classes are given with SD (n=3). Fatty acids Aldehydes Alkanes Primary alcohols Not identified Total  	
    Col-0 5.4 ± 0.48  cer1 9.4±0.75  cer3 1.9±0.13  cer4 5.7±0.45  cer1cer3 4.1±0.32  cer1cer4 6.8±0.47  cer3cer4 1.4±0.09  17.2 ±1.37 74.3 ±5.94 38.8 ± 3.1  18.7±1.26 11.3±0.67 31.3±2.50  6.6±0.45 27.3±2.18 33.3±2.35  22.6±1.80 76.0±6.08 14.9±1.34  3.9±0.35 15.5±1.24 33.5±3.05  13.1±0.92 11.5±0.92 21.3±1.27  6.3±0.44 21.8±1.74 15.9±1.27  6.7 ± 0.53  5.1±0.25  4.5±0.31  6.3±0.44  4.7±0.28  4.4±0.31  4.6±0.37  142.2 ±11.36  71.7 ±6.45  79.8 ± 5.58  120.1 ± 10.81  67.1 ± 5.36  57.4 ± 3.99  56.3 ± 5.04  76	
    200 40 180 160 35  761.2	
  ±	
  68.5  Col-­‐0 cer1  Col-­‐0  140 30  cer1cer3  120 100 20 80 1560  cer4cer1cer3  25  cer1cer4  1040  Fatty	
  a cids  Aldehydes  Alkanes  br-­‐32  br-­‐30  n-­‐34  Primary	
  a lcohols  n-­‐32  n-­‐30  37  35  33  n-­‐28  Alkanes  31  29  27  34  32  30  28  26  24  Aldehydes  n-­‐26  Fatty	
  a cids  34  32  30  28  24 26 28 30 24 26 28 30 25 27 29 31 33 24 26 28 30 32 27 29 31  24  00  Secondary Primary	
   a lcohols alcohols  br-­‐34  520  26  2 Wax	
   coverage	
   [μg/	
   dm Wax	
   coverage	
   [μg	
   dm] 2 ]  	
    	
   	
   Figure 20 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer1, cer3 and cer1cer3 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3.  	
    77	
    775.1	
  ±	
  78.3	
    Col-­‐0 Col-­‐ 0 cer1 cer1  Fatty	
  a cids  Aldehydes  Secondary alcohols Primary	
  a lcohols  br-­‐34  31 br-­‐32  29  br-­‐30  27  n-­‐34  Primary	
  a lcohols  32  n-­‐32  30  n-­‐30  28  n-­‐28  31  Alkanes  26  n-­‐26  29  Alkanes  24  37  33  35  31  33  29 27  27 34  Aldehydes  25 32  30 30  28  28  26  26  24  24  Fatty	
  a cids  30  34  28 30  26 28  24  32  cer4 cer4 cer1cer4 cer1cer4  24  0  761.2	
  ±	
  68.5	
    26  2 Wax	
   coverage	
   [μg/	
   Wax	
   coverage	
   [μg	
  ddm m 2] ]  40 200 180 35 160 30 140 120 25 100 20 80 1560 1040 20 50  	
   	
   Figure 21 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer1, cer4 and cer1cer4 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. 	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    78	
    761.2	
  ±	
  68.5	
    Col-­‐ 0 Col-­‐0  775.1	
  ±	
  78.3	
    cer1 cer3 cer4 cer4  Fatty	
  a cids  Fatty	
  a cids  Aldehydes  Aldehydes  Alkanes  Primary	
  a lcohols  Alkanes  br-­‐34  24 26 28 30 24 26 28 30 25 27 29 31 33 24 26 28 30 32 27 29 31  br-­‐32  br-­‐30  n-­‐34  n-­‐32  n-­‐30  n-­‐28  n-­‐26  37  35  33  31  29  27  34  32  30  28  26  24  34  32  30  28  26  cer1cer4 cer3cer4  24  2 Wax	
   Wax	
  coverage	
   coverage	
  [[μg	
   μg/	
  ddmm 2]]  40  200 35 180 160 30 140 25 120 20 100 80 15 60 10 40 5 20 00  Secondary alcohols  Primary	
  a lcohols  	
   	
   Figure 22 Cuticular wax composition on inflorescence stems of Col-0 (WT), cer3, cer4 and cer3cer4 lines. Levels of major components are expressed as µg/dm2 of stem surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. 	
   	
   	
   	
   	
   	
   	
   	
    	
    79	
    Wax	
  coverage	
  [μg/	
  dm 2 ]  40 Col-­‐0  35  cer1  30  cer3  25  cer1cer3  20 15 10 5  Fatty	
  a cids  Aldehydes  Alkanes  br-­‐34  br-­‐32  br-­‐30  n-­‐34  n-­‐32  n-­‐30  n-­‐28  n-­‐26  37  35  33  31  29  27  34  32  30  28  26  24  34  32  30  28  26  24  0  Primary	
  a lcohols  	
   	
   Figure 23 Cuticular wax composition on rosette leaves of Col-0 (WT), cer1, cer3 and cer1cer3 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. 	
   	
   	
   	
   	
   	
   	
   	
   	
   	
    	
    80	
    Wax	
  coverage	
  [μg/	
  dm 2 ]  40 35  Col-­‐0  30  cer1  25  cer4  20  cer1cer4  15 10 5  Fatty	
  a cids  Aldehydes  Alkanes  br-­‐34  br-­‐32  br-­‐30  n-­‐34  n-­‐32  n-­‐30  n-­‐28  n-­‐26  37  35  33  31  29  27  34  32  30  28  26  24  34  32  30  28  26  24  0  Primary	
  a lcohols  	
   	
   Figure 24 Cuticular wax composition on rosette leaves of Col-0 (WT), cer1, cer4 and cer1cer4 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3. 	
   	
   	
   	
   	
   	
   	
   	
    	
    81	
    Wax	
  coverage	
  [μg/	
  dm 2 ]  40 35  Col-­‐0  30  cer3  25  cer4  20  cer3cer4  15 10 5  Fatty	
  a cids  Aldehydes  Alkanes  br-­‐34  br-­‐32  br-­‐30  n-­‐34  n-­‐32  n-­‐30  n-­‐28  n-­‐26  37  35  33  31  29  27  34  32  30  28  26  24  34  32  30  28  26  24  0  Primary	
  a lcohols  	
   	
   Figure 25 Cuticular wax composition on rosette leaves of Col-0 (WT), cer3, cer4 and cer3cer4 lines. Levels of major components are expressed as µg/dm2 of leaf surface area. Each wax constituent is designated by carbon chain length, and is labeled by chemical class along the x-axis. The data represent mean values ± SD of n=3.  	
    82	
    Chapter 5 Summary and future directions  The cuticle is an asset to the survival of terrestrial plants in several respects. The primary purpose of the cuticle is to create a hydrophobic interface between the plant surface and the aerial environment. Multiple secondary functions also exist as previously introduced (see Section 1.1) Although many details of the wax biosynthetic pathway are well characterized, others need to be further studied and still others confirmed. The objective of this study was to expand the knowledge of the alkane-forming branch of wax biosynthesis in Arabidopsis.   The current work was focused on providing additional evidence challenging the previously described simple two-step pathway hypothesis that includes CER3 as the reductase and CER1 as the decarbonylase. To do this, I have characterized SCD2 and provide evidence for it being involved in the alkane-forming pathway. This is further proof that alkane formation is governed by more than two players.  Besides, a double mutant cer1cer3 was made that can now serve as a tool to test domain functionality of CER1 and CER3 by transforming it with the various constructs of domain truncations and fusions that I previously made. Both these tools will be used in future studies to further elucidate the sequence of enzymatic reactions that form VLC alkanes in Arabidopsis.  The major results of the chapters presented in this thesis will be summarized in the following sections. Further experiments that can build on my results will also be described.       83  5.1	
  Molecular	
  characterization	
  of	
  SCD2	
  and	
  accumulation	
  of	
  aldehydes	
  in	
  leaves	
  of	
   SCD2	
  mutants	
   	
    Several lines of evidence support SCD2’s involvement in the alkane-forming pathway of Arabidopsis leaves and therefore challenging the previously put-forth, simple two-step pathway hypothesis. First, both scd2-1 and scd2-2 leaves had a drastic decrease in alkanes, specifically of C29, C31 and C33, while C30, C32 and C34 aldehydes were elevated. This chemical phenotype was only observed in leaves as stems did not exhibit any change compared to the respective Col7 and Ler ecotypes. The leaf-specific chemical phenotype was in line with SCD2’s highest transcript abundance in leaves and not stems. Furthermore, leaf wax of scd2-1 was restored by transgene complementation. These two lines of evidence confirm that the phenotype was caused by a mutation in the SCD2 gene and not by another gene. Subcellular localization studies in both N. benthamiana and A. thaliana have shown that GFP-tagged SCD2 localizes to the ER, consistent with other characterized genes involved in wax biosynthesis, and providing additional evidence for its role in wax biosynthesis.  Histochemical analysis of a promoter-GUS fusion showed that the SCD2 promoter is active in the phloem of vasculature in two-week-old seedlings. GUS expression was highest in cotyledons, true leaves, roots and shoot apex, while being absent from the hypocotyl and root tips. This data does not correspond with the epidermis-specific site of wax biosynthesis as would be predicted. One reason could be that the 1018 bp promoter region sequence used to construct the pSCD2:GUS was too short so that promoter elements were missing, explaining the localization to the vasculature as an artifact. To remedy this, a longer promoter region of 2000 bp or more can be used for a second construct and compare the histochemical staining with the previous, shorter version.  	
    84	
    While this evidence (apart from promoter-GUS analysis) provides support for SCD2 having a role in alkane formation, it is currently speculative rather than conclusive that it is an enzyme. For this reason, biochemical characterization of SCD2 should be performed in the near future. There should not be obvious obstacles with protein purification, as SCD2’s amino acid sequence does not possess any trans-membrane domains and is predicted to be a soluble protein. Possible future biochemical experiments can be aimed at testing candidate substrates such as VLC aldehydes, especially the C30 molecule synthesized by Chen Peng (M.Sc graduate-Chemistry Department, UBC) first in vivo (S. cerevisiae) and then in vitro. Identification of the C29 alkane as the product would be final proof for SCD2’s function as an enzyme catalyzing the alkaneforming reaction. The precise mechanism of the reaction can further be studied by incorporating the labeled substrate C30 aldehyde (also synthesized by Chen Peng).  To test for the possibility of SCD2 engaging in protein-protein interactions of enzyme complexes involved in wax biosynthesis, two approaches can be used. First, the yeast-two-hybrid assay can be used to test possible interactions between SCD2 and CER1/CER3/CER4/CER8. Alternatively, a BiFC approach can be attempted to transform Arabidopsis mesophyll protoplasts and test the same interactions. While one experiment can be exploratory, the second should be regarded as confirmatory of the potential interactions discovered.  	
    85	
    5.2	
  Characterization	
  of	
  cer1cer3	
  and	
  elucidation	
  of	
  epistatic	
  relationships	
  of	
  CER	
  genes	
   in	
  Arabiopsis	
  wax	
  biosynthesis	
   	
    In Chapter 4, I have generated the cer1cer3 double mutant that has a severe male-sterile phenotype, reduction in wax crystals like the cer3 parent, and has a great decrease in alkane amounts of all chain lengths in both stems and leaves. Most importantly, C30 primary alcohol amounts are slightly lower than in cer3, and much higher than were observed in stems of cer1. This chemical phenotype is the same as the cer3 parent, indicating epistasis. From the epistatic phenotype, it becomes clear that CER3 precedes CER1. This was observed in both stems and leaves. It is noteworthy that chemical analysis on leaves of double cer mutants was never previously reported to the best of my knowledge.  Other double mutants, such as cer1cer4 and cer3cer4 were also generated to study the branchpoint of the wax pathway. No clear epistasis was seen but instead a complex redistribution of constituents from both branches of the pathway was evident. The degree of which depends on the cer mutation at hand. The level of primary alcohols in stems and leaves of cer1cer4 was intermediate to the parents, while in cer3cer4 it was similar to cer4 rather than cer3. This indicates that CER1 and CER3 are genes on the alkane-forming pathway and CER4 on the primary alcohol pathway. In conclusion, this suggests that CER1, CER3 and CER4 encode enzymes that perform modification reactions in both stems and leaves.  Since no compound class was completely reduced in any of the double mutants analyzed in this study, this suggests that there is great redundancy. Triple cer1cer3cer4 or even quadruple mutants cer1cer3cer4cer8 can be made to ascertain redundancy in the alcohol-forming and alkane-forming pathways. These lines can provide further details about the interplay of CER genes in wax biosynthesis. 	
    86	
    The cer1cer3 double mutant is an important tool that can be used to test domain functionality of CER1 and CER3. Previously, I have constructed truncations and fusions between the desaturase and dehydrogenase domains of the two proteins and tested them in C30 fatty acid-producing transgenic yeast. However, no promising results were obtained. In order to continue this research, the double mutant is an ideal in planta background for testing complementation of either or both domains of CER1 and CER3. This will be a long-term project compared to the yeast experiments, due to the time required to obtain Arabidopsis transgenic lines.  	
    87	
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