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Understanding functions of a putative galactose oxidase, RUBY, and its homologues in plant cell wall… Šola, Krešimir 2018

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UNDERSTANDING FUNCTIONS OF A PUTATIVE GALACTOSE OXIDASE, RUBY, AND ITS HOMOLOGUES IN PLANT CELL WALL MODIFICATIONS by  Krešimir Šola  B.Sc., University of Zagreb, 2012  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  December 2018  © Krešimir Šola, 2018  ii  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Understanding Functions of a Putative Galactose Oxidase, RUBY, and Its Homologues in Plant Cell Wall Modifications  submitted by Krešimir Šola in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Botany  Examining Committee: Dr. George Haughn, Botany Supervisor  Dr. Shawn Mansfield, Wood Science Supervisory Committee Member   Supervisory Committee Member Dr. Harry Brumer, Chemistry University Examiner Dr. James Kronstad, Plant Science University Examiner  Additional Supervisory Committee Members: Dr. Ljerka Kunst, Botany Supervisory Committee Member Dr. Xin Li, Botany Supervisory Committee Member iii  Abstract Cell-to-cell adhesion is essential for establishment of multicellularity. In plants, cell adhesion is mediated through a middle lamella composed primarily of pectic polysaccharides, but the molecular interactions that promote and regulate such adhesion are not fully understood. In Chapter 3, Arabidopsis seed coat mucilage was used as a model system to investigate interactions between cell wall carbohydrates. Using a forward-genetic approach, we have discovered a gene encoding a putative galactose oxidase, RUBY PARTICLES IN MUCILAGE (RUBY), that is required for cell-to-cell adhesion in the seed coat epidermis. Cellular and enzymatic analyses support the hypothesis that RUBY facilitates cross-links in the cell walls via the side-chains of rhamnogalacturonan I (RG-I), a constituent of pectin. These results (Chapter 3) provide genetic evidence for oxidative cross-linking in cell walls and assigns a biological function to the galactose/glyoxal oxidase family of enzymes. To better understand functions of galactose oxidases in plants, Arabidopsis homologues of RUBY, GALACTOSE OXIDASE-LIKE (GOXL) genes, were studied (Chapter 4). The expression patterns of these seven genes suggest that all of the members have functions in specialised tissues. Phylogenetic analyses suggest that the GOXL family likely has two pairs of GOXL paralogues, GOXL1 and GOXL6, and RUBY and GOXL3. Surprisingly, RUBY and GOXL3 are expressed in different tissues, whereas GOXL1 and GOXL6 have similar expression patterns, suggesting genetic redundancy. Functional complementation of ruby mutant and qualitative enzyme assays indicate that GOXL1, GOXL3 and GOXL6 are putative galactose oxidases. Plants with mutations in these genes, apart from RUBY, have no obvious phenotypes. When mutations were introduced in both GOXL1 and GOXL6, a collapsed pollen phenotype appeared, indicating that these genes may be redundant. Pollen collapse occurs at the anthesis, when pollen grains are desiccating, suggesting possible roles in pollen wall folding during controlled pollen dehydration (harmomegathy). Further genetic analysis is required to confirm that these mutations are indeed linked to the pollen phenotype. Taken together, putative Arabidopsis galactose oxidases seem to have specialised roles in tissues that may require mechanical support. iv  Lay Summary Cell walls determine the shape of plants and the way in which they grow. They have many functions, including keeping the cells together. I investigated this role of cell walls using a genetic approach. We found that when the product of the RUBY gene does not function, cells separate from one another, suggesting that RUBY is necessary to create strong connections between plant cells. I found that RUBY functions to establish special chemical bonds between molecules of pectin, a gel-like substance abundant in many plant cell walls that is required for cell-cell attachment. Genes similar to RUBY are found in the plant genome. I investigated whether these genes make products with roles analogous to RUBY.  Two such genes, GOXL1 and GOXL6, may work together to prevent collapse of pollen grains. v  Preface This thesis contains 5 chapters, two of which are intended to be published in peer-reviewed journals.  Chapter 1 was written by Krešimir Šola with editorial help from his supervisor, George Haughn. All the figures were made by Krešimir Šola except for figures 1.2 and 1.3 (reprinted with permission).  Chapter 3 and most of Chapter 2 are part of a manuscript submitted as:  Krešimir Šola, Erin J. Gilchrist, David Ropartz, Lisa Wang, Ivo Feussner, Shawn D. Mansfield, Marie-Christine Ralet, George W. Haughn. RUBY, a plant galactose oxidase, promotes pectin cross-links and cell adhesion.  Chapter 2 was written by Krešimir Šola with help from Erin Gilchrist, Marie-Christine Ralet, David Ropartz, Hongwen Chen and Radnaa Naran.  The project described in Chapter 3 was conceived by George Haughn and Erin Gilchrist. Krešimir Šola, George Haughn, Erin Gilchrist and Marie-Christine Ralet designed the research experiments. Erin Gilchrist performed the mutagenesis and isolated ruby mutants. Erin Gilchrist and Lisa Wang positionally cloned the RUBY gene. Erin Gilchrist isolated ruby-4, ruby-5, mum2-10 ruby-5, mum2-1 ruby-4 and ruby-1 bxl1-1 mutants. Marie-Christine Ralet performed sequential extractions of mucilage and analysis of RGase digests. David Ropartz performed IP-RP-UHPLC-MS analyses of RGase for characterisation of branched RG-I and identification of vi  RUBY substrates in mucilage. Monosaccharide linkage (PMAA) analysis, next generation sequencing, and LC-MS analysis of phenolic compounds were analysed as a paid service. Krešimir Šola performed all the remaining experiments and analysed data with supervision from George Haughn. The protein expression experiments in E. coli were performed by Krešimir Šola under the supervision of Ivo Feussner. Monosaccharide composition analyses and RP-HPLC analyses of phenolic compounds were performed by Krešimir Šola under the supervision of Shawn Mansfield. The manuscript draft was written by Krešimir Šola, with editorial help from George Haughn, Marie-Christine Ralet, David Ropartz, Shawn Mansfield, Erin Gilchrist, Ivo Feussner, and Lisa Wang. All the figures were made by Krešimir Šola except for figure 3.6 C and D, and figure 3.9 E to H, which were made by David Ropartz.  Research outlined in Chapter 4 was conceived and experiments designed by George Haughn and Krešimir Šola. Erin Gilchrist isolated goxl2-1 and goxl3-1 lines. Mahsa Movahedan, Julia Lohmann and Yi Li performed experiments and analysed data under the supervision of Krešimir Šola. Mahsa Movahedan, Julia Lohmann, Yi Li and Krešimir Šola isolated the remaining insertional mutant lines. Mahsa Movahedan and Julia Lohmann constructed promoter-GUS plasmids. Yi Li and Krešimir Šola constructed ProRUBY:GOXL-Citrine plasmids. Julia Lohmann, Yi Li and Krešimir Šola performed histochemical GUS experiments. The remaining experiments were performed, and data analysed, by Krešimir Šola. Krešimir Šola wrote the chapter with editorial help from George Haughn. All the figures were made by Krešimir Šola, except for figure 4.1 (reprinted with permission).  Chapter 5 was written by Krešimir Šola with editorial help from George Haughn. vii  Table of Contents  Abstract ......................................................................................................................................... iii Lay Summary ............................................................................................................................... iv Preface .............................................................................................................................................v Table of Contents ........................................................................................................................ vii List of Tables ............................................................................................................................... xii List of Figures ............................................................................................................................. xiii List of Abbreviations ................................................................................................................. xvi Acknowledgements ......................................................................................................................xx Dedication .................................................................................................................................. xxii Chapter 1: Introduction ................................................................................................................1 1.1 Composition and organisation of type I plant cell walls................................................. 1 1.1.1 Cellulose ..................................................................................................................... 4 1.1.2 Hemicelluloses ............................................................................................................ 5 1.1.2.1 Xyloglucan .......................................................................................................... 5 1.1.2.2 Xylans ................................................................................................................. 6 1.1.2.3 Mannans .............................................................................................................. 7 1.1.3 Pectins ......................................................................................................................... 7 1.1.3.1 Homogalacturonan .............................................................................................. 8 1.1.3.2 Rhamnogalacturonan-I ........................................................................................ 9 1.1.3.3 Rhamnogalacturonan-II .................................................................................... 10 1.1.4 Hydroxyproline-rich glycoproteins ........................................................................... 13 viii  1.1.4.1 Extensins ........................................................................................................... 14 1.1.4.2 Arabinogalactan proteins .................................................................................. 15 1.1.5 Callose....................................................................................................................... 16 1.1.6 Primary cell wall assembly, structure, and organisation ........................................... 17 1.2 Arabidopsis seed coat mucilage .................................................................................... 22 1.2.1 Mucilage is produced from a single cell type ........................................................... 23 1.2.2 Composition of the seed coat mucilage .................................................................... 24 1.2.3 Structure of the seed coat mucilage .......................................................................... 27 1.2.4 Function of cell wall components in the mucilage ................................................... 27 1.2.5 Removal of RG-I side-chains is needed for extrusion .............................................. 29 1.3 Research questions and goals........................................................................................ 30 Chapter 2: Materials and Methods ............................................................................................32 2.1 Plant materials and growth conditions .......................................................................... 32 2.1.1 Arabidopsis thaliana ................................................................................................. 32 2.2 Statistical Analyses ....................................................................................................... 34 2.3 Microscopy ................................................................................................................... 35 2.3.1 Light microscopy ...................................................................................................... 35 2.3.2 Spinning disk confocal microscopy .......................................................................... 36 2.3.3 Scanning electron microscopy (SEM) ...................................................................... 37 2.4 Genotyping .................................................................................................................... 37 2.5 Mutagenesis, genetic analysis, and positional cloning of ruby ..................................... 40 2.6 Molecular cloning and transgenic plants ...................................................................... 45 2.6.1 ProRUBY:RUBY-Citrine........................................................................................... 45 ix  2.6.2 ProRUBY:GOXL-Citrine .......................................................................................... 47 2.6.3 ProGOXL:GUS ......................................................................................................... 48 2.6.4 CRISPR-Cas9 mutagenesis ....................................................................................... 49 2.7 Gene expression analyses ............................................................................................. 50 2.7.1 Reverse-transcription PCR (RT-PCR) ...................................................................... 50 2.7.2 Histochemical promoter-GUS assays ....................................................................... 51 2.8 Chemical analyses ......................................................................................................... 52 2.8.1 Monosaccharide composition analysis using HPAEC-PAD .................................... 52 2.8.2 Per-O-methylation and Linkage analysis of neutral sugars ...................................... 55 2.8.3 Digestion and analysis of mucilage RG-I ................................................................. 55 2.8.4 IP-RP-UHPLC-MS/MS ............................................................................................ 56 2.8.5 Extraction and analysis of seed surface phenolics .................................................... 57 2.9 Phylogenetic analysis .................................................................................................... 58 2.10 Protein expression and purification .............................................................................. 59 2.10.1 Protein expression and purification in E. coli ....................................................... 59 2.11 Enzyme activity experiments ........................................................................................ 61 2.11.1 Enzyme assays with seeds .................................................................................... 61 2.11.2 Detection of galactose oxidase substrates in the mucilage ................................... 62 2.11.3 Mucilage insolubility experiments ........................................................................ 63 Chapter 3: RUBY PARTICLES IN MUCILAGE (RUBY) is a putative galactose oxidase that cross-links pectin ..................................................................................................................65 3.1 Introduction ................................................................................................................... 65 3.2 Results ........................................................................................................................... 67 x  3.2.1 Mutations in RUBY can suppress the mum2 mucilage extrusion phenotype ............ 67 3.2.2 RUBY is required for mucilage integrity.................................................................. 70 3.2.3 RUBY is required for cell-to-cell adhesion .............................................................. 73 3.2.4 RUBY is needed to cross-link arabinogalactan-branched RG-I to the seed ............. 75 3.2.5 RUBY encodes a putative galactose oxidase ............................................................. 82 3.2.6 RUBY is expressed in seeds after mucilage secretion and localises to the apoplast in columella ............................................................................................................................... 94 3.2.7 Pectin cross-linking is most likely mediated through hemiacetals, not through hydroxycinnamate esters ....................................................................................................... 98 3.3 Discussion ................................................................................................................... 104 3.3.1 RUBY is a putative galactose oxidase with RG-I side-chains as its substrate in the mucilage .............................................................................................................................. 104 3.3.2 Oxidation of galactose most likely creates hemiacetal bonds in the mucilage ....... 106 3.3.3 Mutations in RUBY affect mucilage appearance .................................................... 109 Chapter 4: Functional analysis of the GALACTOSE OXIDASE-LIKE (GOXL) gene family from Arabidopsis thaliana ..........................................................................................................111 4.1 Introduction ................................................................................................................. 111 4.1.1 Pollen development ................................................................................................. 111 4.1.2 Pollen wall structure and composition .................................................................... 114 4.1.3 Objectives ............................................................................................................... 116 4.2 Results ......................................................................................................................... 117 4.2.1 Phylogenetic analysis of GOXL proteins from Arabidopsis .................................. 117 4.2.2 Tissue-wide expression analysis of GOXL genes ................................................... 121 xi  4.2.3 Functional complementation of ruby with GOXL genes ........................................ 131 4.2.4 Reverse-genetic analysis of GOXL mutants ........................................................... 136 4.3 Discussion ................................................................................................................... 140 4.3.1 Some GOXL genes have undergone duplications and regulatory neofunctionalisation or subfunctionalisation ........................................................................................................ 140 4.3.2 GOXLs likely have distinct biological functions .................................................... 141 4.3.2.1 GOXL2 ........................................................................................................... 141 4.3.2.2 GOXL3 ........................................................................................................... 142 4.3.2.3 GOXL4 ........................................................................................................... 143 4.3.2.4 GOXL5 ........................................................................................................... 143 4.3.2.5 GOXL1 and GOXL6 possibly function to establish proper pollen wall structure 144 Chapter 5: Conclusions .............................................................................................................147 5.1 Galactose oxidase can cross-link plant cell walls ....................................................... 147 5.1.1 Future directions ..................................................................................................... 147 5.2 Putative galactose oxidases in Arabidopsis may function in specialised tissues ........ 148 5.2.1 Future directions ..................................................................................................... 151 5.3 Summary ..................................................................................................................... 153 Bibliography ...............................................................................................................................154  xii  List of Tables Table 2.1: A list of mutant lines used in this study ....................................................................... 32 Table 2.2: A list of primers used to genotype mutations used in this study ................................. 37 Table 2.3: A list of primers used in crude-mapping ..................................................................... 43 Table 2.4: A list of primers used to clone GOXL genes in-frame with Citrine ............................ 48 Table 2.5: A list of primers used to amplify putative promoter regions of GOXL genes ............. 49 Table 2.6: A list of primers used in RT-PCR reactions ................................................................ 51 Table 3.1: Monosaccharide linkage analysis of wild-type and ruby-1 mucilage ......................... 80 Table 3.2: A list of compounds tested as substrates for oxidases on dry mature seeds using HRP-TMB assay .................................................................................................................................... 89 Table 4.1: Arabidopsis thaliana members of GALACTOSE OXIDASE-LIKE (GOXL) family of proteins with predicted subcellular location and signal peptide based on Aramemnon database..................................................................................................................................................... 121 Table 4.2: Results of functional complementation of ruby by GOXL genes based on ruthenium red staining after hydration in water ........................................................................................... 133 xiii  List of Figures Figure 1.1: Generalised structures of most components of type I primary cell walls .................... 3 Figure 1.2: Structure and cross-linking of RG-II .......................................................................... 12 Figure 1.3: Schematic representation of typical O-glycosylation patters of plant HRGPs .......... 14 Figure 1.4: Oxidative cross-linking of HRGPs through tyrosines (Tyr) ...................................... 19 Figure 1.5: Structures of dimers and trimers of hydroxycinnamic acids isolated from plant cell walls .............................................................................................................................................. 21 Figure 1.6: Hydrated wild-type seeds (Col-2) of Arabidopsis thaliana stained with Ruthenium Red ................................................................................................................................................ 22 Figure 1.7: Developmental sequence of a seed coat epidermal cell ............................................. 24 Figure 1.8: Polysaccharide components of the Arabidopsis seed coat mucilage ......................... 26 Figure 1.9: Phenotypes of mucilage-modified2 (mum2) ............................................................... 30 Figure 3.1: ruby exhibits multiple seed coat mucilage phenotypes .............................................. 69 Figure 3.2: ruby suppresses mum2 and bxl1 phenotypes to a different degree............................. 72 Figure 3.3: RUBY is involved in cell-cell adhesion between seed coat epidermal cells as well as between seed coat epidermal cells and palisade cells ................................................................... 74 Figure 3.4 Cell adhesion defects of ruby-1 are enhanced by chelator (EDTA) and suppressed by calcium (CaCl2) ............................................................................................................................. 75 Figure 3.5: Branched RG-I is present in ruby mucilage ............................................................... 77 Figure 3.6: RGase releases novel branched RG-I from ruby-1 mucilage ..................................... 81 Figure 3.7: RUBY encodes a glyoxal oxidase-like protein .......................................................... 83 Figure 3.8: Insertional mutant, ruby-5, resembles ruby-1 in all phenotypic aspects .................... 85 Figure 3.9: RUBY is a putative galactose oxidase functioning on mum2 mucilage..................... 87 xiv  Figure 3.10: RUBY contains catalytic amino acids required for galactose/glyoxal oxidase function ......................................................................................................................................... 92 Figure 3.11: Western blot analysis of purification of RUBY-His6 expressed in Arctic Express (DE3) strain of E. coli ................................................................................................................... 93 Figure 3.12: RUBY-Citrine is expressed after mucilage secretion and localises to the apoplast . 96 Figure 3.13: RUBY is expressed in root epidermis and seed coat................................................ 97 Figure 3.14: RUBY most likely functions through hemiacetal formation, not through hydroxycinnamate cross-linking ................................................................................................... 99 Figure 3.15: Most abundant phenolics on seed surface are sinapic acid and a flavonol glycoside..................................................................................................................................................... 102 Figure 3.16: A model proposing cross-linking of branched RG-I by galactose oxidation ......... 103 Figure 4.1: Schematic representation of transverse section through an anther during pollen development ................................................................................................................................ 113 Figure 4.2: Development of the pollen wall ............................................................................... 114 Figure 4.3: Pollen wall structure of the mature pollen grain ...................................................... 116 Figure 4.4: Unrooted Maximum Likelihood phylogenetic tree of GOXL proteins from Arabidopsis thaliana compared with putative glyoxal oxidase from Vitis pseudoreticulata (VpGLOX) and characterised fungal galactose oxidases, alcohol oxidases and glyoxal oxidases (CAZy AA5 family). ................................................................................................................... 118 Figure 4.5: Unrooted Maximum Likelihood phylogenetic tree of angiosperm GOXL proteins 120 Figure 4.6: The structure of GOXL genes including the entire transcriptional regulatory region between the GOXL 5ʹ UTR and the nearest gene upstream ........................................................ 122 xv  Figure 4.7: ProGOXL1 and ProGOXL6 are transcriptionally active late in pollen development..................................................................................................................................................... 124 Figure 4.8: ProGOXL2 is active in the developing pistil, silique septum and funiculi .............. 126 Figure 4.9: ProGOXL3 is active in the phloem .......................................................................... 128 Figure 4.10: ProGOXL4 is active in the tapetum of developing anthers .................................... 129 Figure 4.11: ProGOXL5 is active in the parenchyma ................................................................. 130 Figure 4.12: T2 seeds of ProRUBY:GOXL-Citrine in mum2-1 ruby-1 plants ............................ 131 Figure 4.13: HRP-TMB colourimetric assay demonstrating that galactose oxidase activity is restored in ProRUBY:GOXL-Citrine lines .................................................................................. 135 Figure 4.14: Subcellular distribution of GOXL-Citrine in the seed coat epidermis of seeds from senescing siliques ........................................................................................................................ 136 Figure 4.15: The line carrying goxl1-4 and goxl6-5 mutations is partially male sterile due to collapsed pollen grains ................................................................................................................ 138 Figure 4.16: Phenotype of the putative mutant becomes apparent at the anthesis ..................... 139 Figure 4.17: Proposed harmomegathy-based model showing collapse of the pollen grain during desiccation in the putative goxl1 goxl6 mutant ........................................................................... 146 xvi  List of Abbreviations 4CL 4-COUMARATE:CoA LIGASE AcOH Acetic acid ACT2 ACTIN2 AGP Arabinoagalactan protein AIR Alcohol-insoluble residue ANOVA Analysis of variance AOMT1 CAFFEOYL COENZYME A O-METHYLTRANSFERASE1 Api Apiose Ara Arabinose AX Arabinoxylan BXL1 BETA-XYLOSIDASE1 CESA Cellulose synthase A COMT1 CAFFEOYL-O-METHYLTRANSFERASE1 CRISPR Clustered-regularly interspaced short palindromic repeats CSLA2 CELLULOSE SYNTHASE-LIKE A2 di-IDT Di-isodityrosine DM Degree of methylesterification DPA Days post-anthesis EMS Ethyl methanesulphonate EtOAc Ethyl acetate EtOH Ethanol EXT Extensin xvii  FA Ferulic acid FAH1 FERULIC ACID 5-HYDROXYLASE1 FLA Fasciclin-like arabinogalactan protein FLY1 FLYING SAUCER1 FTA Flinders Technology Associates Fuc Fucose Gal Galactose GalOx Galactose oxidase GalA Galacturonic acid GaM Galactomannan GAPC1 GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE C SUBUNIT1 GAX Glucuronoarabinoxylan GGM Galactoglucomannan Glc Glucose GlcUA Glucuronic acid GOXL GALACTOSE OXIDASE-LIKE GM Glucomannan GPI Glycosylphosphatidylinositol GX Glucuronoxylan H2O2 Hydrogen peroxide H2SO4 Sulphuric acid HCl Hydrochloric acid xviii  HG Homogalacturonan HPAEC High-performance anion exchange chromatography HPLC High-performance liquid chromatography HRGP Hydroxyproline-rich glycoprotein HRP Horseradish peroxidase IDT Isodityrosine IRX IRREGULAR XYLEM KHCO3 Potassium hydrogencarbonate Man Mannose MeCN Acetonitrile MeOH Methanol MS Mass spectrometry MUCI MUCILAGE-RELATED MUM2 MUCILAGE-MODIFIED2 Na2CO3 Sodium carbonate NaBH4 Sodium borohydride NaOAc Sodium acetate NaPi Sodium phosphate NMR Nuclear magnetic resonance PAD Pulsed amperometric detection PCD Programmed cell death PME PECTIN METHYLESTERASE PMEI PECTIN METHYLESTERASE INHIBITOR xix  REF1 REDUCED EPIDERMAL FLUORESCENCE1 RG-I Rhamnogalacturonan-I RG-II Rhamnogalacturonan-II Rha Rhamnose RUBY RUBY PARTICLES IN MUCILAGE SA Sinapic acid SD Standard deviation SEM Scanning electron microscopy SOS5 SALT-OVERLY SENSITIVE5 T-DNA Transfer DNA TEM Transmission electron microscopy TFA Trifluoroacetic acid TMB 3, 3ʹ, 5, 5ʹ - Tetramethylbenzidine XyG Xyloglucan Xyl Xylose  xx  Acknowledgements First and foremost, I would like to thank Dr. George Haughn for accepting me as his student and for providing mentorship in many aspects of my academic career. I am especially grateful to him for giving me the opportunity to delve deeper into areas that are usually not within his laboratory’s expertise, and for showing patience and understanding when many experiments turned out to be unsuccessful. I am very grateful to all my supervisory committee members. I thank Dr. Ljerka Kunst for encouraging me to be more critical, for providing numerous suggestions on how to improve my experiments and presentations, as well as career advice. I would also like to thank Dr. Shawn Mansfield for providing me with the opportunity to perform analytical experiments in his laboratory and for his advice on experimental design and data analysis. I am thankful to Dr. Xin Li for her feedback, suggestions, and a unique perspective on my project. I am very grateful to Dr. Ivo Feussner (University of Goettingen) for welcoming me into his laboratory for two months. His kindness, encouragement, and advice came at the time when I needed them most. Many thanks to all the members of Haughn, Kunst, Feussner and Mansfield labs. It was wonderful to work in such supportive, stimulating, and diverse environments. My greatest thanks go to Dr. Erin Gilchrist who proved to be the best mentor I could ask for. Her patience, understanding, encouragement and support helped me develop as a scientist. Many thanks to Dr. Gillian Dean who had helped me immensely in many ways. Her advice, critique, and encouragement were invaluable. I am very grateful to have had wonderful mentees, Mahsa Movahedan, Julia Lohmann, and Yi Li, who helped me complete work on RUBY homologues. Thanks to all the other members of Haughn and Kunst labs, especially Robert McGee, Dr. Yi-xxi  Chen Lee, Dr. Tegan Haslam, Dr. Shuang Liu, Dr. Lifang Zhao, Dr. Gabriel Levesque-Tremblay, and Dr. Jonathan Griffiths who helped me numerous times and shared many academic ups and downs with me. I am very grateful to the members of the Mansfield lab, especially Dr. Faride Unda for many wonderful discussions, her help with experimental design and troubleshooting, and taking care of my samples when I was not able to. I would also like to thank Dr. Eliana Gonzales-Vigil, Pablo Antonio Chung, Yaseen Mottiar, and Dr. Letitia Da Ros who helped me through discussions and saved my experiments from failure. I am grateful to all the members of the Feussner lab, with special thanks to Katharina Vollheyde and Dr. Ellen Hornung for being amazing hosts. Many thanks to our collaborators, Dr. Marie-Christine Ralet and Dr. David Ropartz for doing wonderful work, as well as Marie-Jeanne Crépeau and Jacqueline Vigouroux who assisted them. Thanks to Hongwen Chen (Mass Spectrometry Services, Department of Chemistry, Simon Fraser University) for performing LC-ESI-MS analysis; Radnaa Naran (Complex Carbohydrate Research Center, University of Georgia) for performing PMAA linkage analysis; Dr. Yunchen Gong (Centre for the Analysis of Genome Evolution and Function, University of Toronto) for performing Next Generation Sequencing and data processing; UBC Bioimaging Facility, especially Derrick Horne and Kevin Hodgson; Dr. Yann Mathieu and Dr. Harry Brumer for help with my Pichia pastoris expression experiments and advice on enzyme assays. I would like to thank the Department of Botany and the University of British Columbia for supporting me through a Four-Year Fellowship, and IRTG 2172 "PRoTECT" grant from the German Research Foundation (DFG) for supporting my visit to the Feussner lab. I am grateful to have had lots of support and encouragement from my friends and family, without whom I may not have been able to complete my studies. xxii  Dedication  To my grandmother, Vesna Weiss-Rogoz.1  Chapter 1: Introduction Almost all plant cells are surrounded by an extracellular matrix called the cell wall that acts as an interface between other cells and the external environment. Cell walls provide mechanical support, act as a physical barrier to pests and pathogens, mediate cell-cell adhesion, determine cell shape, and serve as a medium for cell-cell communication. The initial or primary cell wall, established at the time of cell division, is most commonly composed primarily of several  classes of polysaccharides: pectins, cellulose and hemicelluloses (Cosgrove, 2005) although the specific composition and arrangement of components varies greatly among cell types and species. The typical primary cell wall must be relatively plastic to accommodate growth and expansion. Additional specialised cell wall material, termed the secondary cell wall, may be added between the plasma membrane and primary cell wall in some differentiating cell types. Secondary cell wall composition of different cell types is even more variable than that of the primary wall. Although the general nature of cell walls has been established, there remain large gaps in our understanding of the structure-function relationships. The main focus of my research is on the cell wall carbohydrate class pectin. More specifically, I investigated how modifications to the structure of the pectin component rhamnogalacturonan-I can impact both its physical properties and its biological function. By way of background, below I review the current state of knowledge of the composition and structure of cell walls in general, as well as that of seed mucilage.  1.1 Composition and organisation of type I plant cell walls Type I primary cell walls have been described by Carpita and Gibeaut (1993) as generalised cell walls of most flowering plants. Its main components are cellulose microfibrils, hemicelluloses, mainly xyloglucan, structural proteins, and pectins. In this model, cellulose microfibrils are 2  cross-linked by xyloglucan into a single network, which is embedded in a pectin matrix. While the described composition of type I cell walls is accurate, recent advancements demonstrated that matrix polysaccharides, hemicelluloses and pectins, as well as structural proteins, can be covalently linked to one another (Tan et al., 2013). Type I and Type II cell walls (eg. those of Poales) differ in their matrix carbohydrate composition where pectins predominate in Type I cell walls while xylans are more abundant in type II cell walls (Carpita and Gibeaut, 1993). Since my research focuses on Type I cell walls, I have reviewed their composition below.                3   Figure 1.1: Generalised structures of most components of type I primary cell walls. Figure based on Scheller et al., 2006 and Scheller and Ulvskov, 2010.  4  1.1.1 Cellulose Cellulose is the most abundant cell wall polysaccharide, found in both primary and secondary cell walls. It is made of D-glucopyranose (D-Glcp) arranged into linear chains by β-(1→4) glycosidic bonds (Figure 1.1). The number of glucose molecules in a single chain varies from  500 to 14000 units (Somerville, 2006). These chains are synthesised by CELLULOSE SYNTHASE A (CESA) proteins, which form rosette-shaped cellulose synthase complexes (CSCs) in the plasma membrane. CSCs are comprised of six subunits, each of which is believed to contain three distinct CESA isoforms (Vandavasi et al., 2016; Hill et al., 2014). Most plant species contain a family of related CESA proteins, and different combinations of CESAs are used to make distinct primary and secondary cell walls. For example, in Arabidopsis, primary cell wall cellulose is synthesised by CESA1, CESA3 and one of CESA2, CESA5, CESA6 or CESA9, which are redundant and tissue-specific (Mendu et al., 2011; Persson et al., 2007; Desprez et al., 2007). In contrast, lignified secondary walls, rich in cellulose, are made by complexes comprising CESA4, CESA7 and CESA8 (Gardiner et al., 2003; Atanassov et al., 2009; Taylor et al., 2003). Multiple individual chains of cellulose are synthesised simultaneously by each CESA polypeptide of the complex, which then interact with each other through hydrogen bonds to form microfibrils. Recent studies indicate that some functional CESA complexes are composed of a single CESA isoform (Cho et al., 2017; Purushotham et al., 2016) indicating that a CESA heterocomplex is not necessary for formation of microfibrils.  Cellulose microfibrils serve as the load bearing component of the cell wall (Willats et al., 2001a). Their deposition can determine the way in which cells elongate by resisting the turgor-driven expansion in one or more planes (Roelofsen, 1966). If microfibrils are deposited parallel to one another, the cell will be able to expand only perpendicular to the microfibrils, resulting in 5  elongation in a single plane. On the other hand, deposition of microfibrils in random directions will result in isometric cell expansion, giving the cell a spherical shape.  1.1.2 Hemicelluloses Cellulose microfibrils are cross-linked by matrix polysaccharides – hemicelluloses and pectins to form a strong extracellular network. Hemicelluloses are glycans with backbones containing β-(1→4)-linked D-Glcp, D-xylopyranose (D-Xylp), or D-mannopyranose (D-Manp), with glycosidic bond in equatorial orientation (Scheller and Ulvskov, 2010).   1.1.2.1 Xyloglucan The most abundant hemicellulose in primary cell walls of seed plants (except grasses) is xyloglucan (XyG; Scheller and Ulvskov, 2010). This polysaccharide contains (1→4)-β-D-Glcp backbone and side-chains with specific nomenclature depending on their composition (Fry et al., 1993; Hayashi T., 1989; Scheller and Ulvskov, 2010). Individual glucoses in XyG can be named G for a unit of Glc without a side-chain, X for Glc with a single D-Xylp as a side-chain, L for D-galactopyranose (D-Galp)-D-Xylp side-chain, and F for L-fucopyranose (L-Fucp)-D-Galp-D-Xylp, representing the most common configurations of side-chains in Eudicots (Figure 1.1; Hayashi T., 1989; Fry et al., 1993). XyG is deposited in the growing cell wall starting with cell division, and remains in the cell wall during cell growth (Moore and Staehelin, 1988; Hayashi T., 1989). It associates with the surface of cellulose microfibrils through hydrogen bonding to form cross-links between them (Dick-Pérez et al., 2011; Zheng et al., 2018; Whitney et al., 1995). XyG does not appear to be essential since Arabidopsis mutants lacking xyloglucan are viable 6  (Cavalier et al., 2008).  However, it does have roles in regulation of cell wall stiffness and strength (Cavalier et al., 2008; Zabotina et al., 2012).  1.1.2.2 Xylans Xylans are the most abundant hemicelluloses, commonly associated with secondary cell walls in Eudicots (Ebringerová and Heinze, 2000; Scheller and Ulvskov, 2010), but they have also been detected in primary cell walls (Darvill et al., 1980). Xylans of seed plants contain (1→4)-β-D-Xylp backbone, with most common types of single monosaccharide branches being D-glucopyranosyluronic acid (D-GlcpUA) or 4-O-methyl-GlcpUA linked by an α-(1→2) bond to the Xyl in the backbone (glucuronoxylans; GX; Figure 1.1), and α-L-arabinofuranose (L-Araf) linked to Xyl in the backbone at C-2, or both C-2 and C-3 positions (arabinoxylans; AX; Figure 1.1).  Glucuronoarabinoxylans (GAX) come with both GlcUA and Ara side-chains (Ebringerová and Heinze, 2000). Xylose in the backbone is frequently acetylated at C-2 and/or C-3 positions in the pyranose ring (Kabel et al., 2003; Busse-Wicher et al., 2016), whereas Ara in AX in grasses often contain ferulic acid linked through an ester bond at the C-5 in the furanose ring (de O. Buanafina, 2009; Wende and Fry, 1997). Xylans bind to cellulose microfibrils through hydrogen bonds, where lower degrees of branching result in better binding (Köhnke et al., 2011; Kabel et al., 2007). Upon binding to cellulose, xylan changes conformation to assume a similar conformation to that of cellulose (Simmons et al., 2016; Busse-Wicher et al., 2016). Based on dwarf phenotypes of O-acetyltransferase mutants of Arabidopsis, acetylation of xylans appears to be required for normal growth (Yuan et al., 2016; Xiong et al., 2013). The removal of acetyl groups from xylan results in changes in the branching patterns, reduced binding of xylan to cellulose microfibrils, and altered mechanical properties of cell walls (Grantham et al., 2017). 7  The association of xylan with cellulose may be disrupted in O-acetyltransferase mutants because alterations in the spacing of side-chains result in a xylan with a stereochemistry that is incompatible for interacting with cellulose  (Busse-Wicher et al., 2016).  1.1.2.3 Mannans Mannans are hemicelluloses with backbones containing D-mannopyranose (D-Manp; mannans) or both D-Manp and D-Glcp (glucomannans; GM) linked through β-(1→4) glycosidic bonds (Ebringerova et al., 2005). They usually have single α-(1→6)-linked D-Galp residues as branches, resulting in polysaccharides named galactomannans (GaM; Figure 1.1). GM in primary walls was found to be branched, therefore named galactoglucomannans (GGM; Figure 1.1), whereas branching was low in the secondary cell walls (Ebringerova et al., 2005). Mannans are able to self-aggregate and form crystalline lattices, which is negatively correlated with branching (Kapoor et al., 1995, 1998; Ebringerova et al., 2005). They can associate with cellulose microfibrils and cross-link them to form a network. Similar to self-aggregation, lower branching results in stronger association (Whitney et al., 1998). Mannans are often acetylated, which, similar to Gal branching, prevents aggregation (Ebringerova et al., 2005). Mutants deficient in mannan biosynthesis appear to have defects in embryo (Goubet et al., 2003, 2009), root hair and vascular development (Yin et al., 2011), indicating that these polysaccharides have important roles in plant development.   1.1.3 Pectins Acidic polysaccharides that contain (1→4)-linked α-D-galactopyranosyluronic acid (D-GalpA) in their backbone are called pectins (Ridley et al., 2001). They are highly abundant in primary 8  cell walls of gymnosperms, Eudicots and monocots other than Poales (Ridley et al., 2001; Mohnen, 2008). There are three major types of pectins with structural differences that result in unique properties and functions. Evidence suggests that all three types of pectin are covalently linked to one another, and can therefore be considered as three different domains of a single molecule (Ridley et al., 2001; Willats et al., 2001a). Together they form a strong hydrophilic carbohydrate network that is reinforced by cellulose microfibrils. Pectins are enriched in the area of the extracellular matrix shared by two adjacent cells, the middle lamella, and junctions where more than two cells meet, suggesting a role in cell adhesion (Willats et al., 2001a; Mohnen, 2008).  1.1.3.1 Homogalacturonan The simplest and most abundant pectin domain is homogalacturonan (HG), being a α-(1→4)-linked galacturonan without side-chains (Figure 1.1). Carboxylic groups at C-6 in GalA residues can be free or methyl-esterified, which can dynamically change the properties of this polysaccharide (Ridley et al., 2001). HG is synthesised in highly esterified form, but it can be de-esterified by pectin methylesterases (PMEs) in the cell wall. Once carboxyl groups are de-esterified by PMEs, they can bind Ca2+ and form cross-links between two HG chains, depending on the pattern of de-esterification (Pelloux et al., 2007; Levesque-Tremblay et al., 2015). De-esterification can be blockwise or non-blockwise (Kohn et al., 1983). When PMEs de-esterify sequential GalAs (blockwise), the negatively charged stretches of carboxyl groups can interact with Ca2+ to form calcium bridges that cross-link HG (Limberg et al., 2000). On the other hand, random de-esterification by PMEs (non-blockwise) leads to an HG molecule that is prone to endo-polygalacturonase degradation (Limberg et al., 2000). GalAs from HG were also found to 9  be O-acetylated to varying degrees at the C-2 and C-3 positions (Ishii, 1997). HG has many important biological functions. Its dynamic cross-linking is utilised in regulation of the cell wall stiffness, as demonstrated by roles in maintaining mechanical support (Hongo et al., 2012), cell adhesion (Orfila et al., 2005; Bouton et al., 2002; Atkinson et al., 2002), cell separation (Francis et al., 2006; Rhee and Somerville, 1998; Rhee et al., 2003), pollen germination (Leroux et al., 2015), and pollen tube growth (Jiang et al., 2005). Products of HG degradation, oligogalacturonides, are used as signalling molecules in response to pathogen attack (Ferrari et al., 2013) or as a developmental cue (Sinclair et al., 2017).  1.1.3.2 Rhamnogalacturonan-I The most structurally diverse pectin domain is rhamnogalacturonan-I (RG-I). Unlike other pectins, it has a backbone of alternating D-GalpA and L-rhamnopyranose (L-Rhap) in a [→2)-α-L-Rhap-(1→4)-α-D-GalpA-(1→] pattern (Stevenson et al., 1980; Lau et al., 1985). Diversity of branching is what makes RG-I complex. It typically branches at the C-4 of Rha, even though exceptions have been described (Naran et al., 2008). These branches usually contain exclusively D-Galp (galactans), exclusively L-Araf (arabinans), or both (arabinogalactans) (Figure 1.1; Lau et al., 1987; Lerouge et al., 1993). Chain lengths and linkages usually depend on cell types and developmental stages (McCartney et al., 2000; Lee et al., 2013). Arabinans usually contain α-(1→5)-linked chains that can branch at the C-2 and C-3 of furanose rings. Galactans most commonly appear as linear β-(1→4)-linked chains, as well as 3- and 6-linked β-D-galactans (Albersheim et al., 2011; Ridley et al., 2001). Like HG, RG-I can also be O-acetylated at the C-2 and/or C-3 position of GalpA (Ishii, 1997; Komalavilas and Mort, 1989). Ferulic acid esters (ferulates) and their dimers have been found attached to arabinan and galactan side-chains of 10  RG-I, suggesting that they are involved in side-chain cross-linking (Ralet et al., 2005). Similar to HG and consistent with pectin being a single molecule, RG-I has roles in regulation of mechanical properties (Verhertbruggen et al., 2013) and cell adhesion (Molina-Hidalgo et al., 2013; Stolle-Smits et al., 1999; Redgwell et al., 1992; Iwai et al., 2001). This is likely facilitated through hydrophobic interactions and hydrogen bonds between longer, linear β-(1→4)-galactans (Makshakova et al., 2017), but the roles of shorter side-chains or side-chains of different composition or linkages still remain elusive.  1.1.3.3 Rhamnogalacturonan-II Compositionally, the most complex pectic polysaccharide is rhamnogalacturonan-II (RG-II). It contains a backbone that is identical to HG, and side-chains composed of 12 different monosaccharides with more than 20 distinct linkages in a structurally conserved order (O’Neill et al., 2004). Four separate side-chains are attached to the HG-like backbone: sidechains A and B through the C-2, and side-chains C and D through the C-3 of GalA residue (Figure 1.2 A; O’Neill et al., 2004). Chains C and D contain 2-keto-3-deoxy-D-manno-octulosonic acid (Kdo) and 2-keto-3-deoxy-D-lyxo-heptulosaric acid (Dha), respectively, which appear in plants only as a part of RG-II (Albersheim et al., 2011). Aside from D-GalpA, L-Araf, D-Galp, L-Rhap, D-glucopyranosyluronic acid (D-GlcpUA) and L-fucopyranose (L-Fucp), monosaccharides commonly found in cell wall polysaccharides, rare monosaccharides such as D-apiose (D-Apif), L-aceric acid (L-AcefA), 2-O-methyl L-fucose (2Me-L-Fuc), 2-O-methyl D-xylose (2Me-D-Xyl), and L-galactose (L-Gal) comprise side-chains of RG-II (Figure 1.2 A; O’Neill et al., 2004). Unlike HG, which cross-links through Ca2+ bridges, two monomeric RG-II (mRG-II) molecules can dimerise to form dRG-II through borate diesters formed between L-Apif residues of A chains 11  (Figure 1.2 B and C; Ishii et al., 1999). This cross-linking happens in the endomembrane system prior to the deposition of RG-II in the cell wall (Chormova et al., 2014). Even though L-Apif is the first residue in chain A, borate cross-linking requires the entire chain structure (O’Neill et al., 2001; Iwai et al., 2002). Despite the low abundance of RG-II in cell walls, the lack of borate cross-links results in stunted growth and loss of cell adhesion, suggesting that this polymer has important roles in determining cell wall properties (O’Neill et al., 2001; Iwai et al., 2002). 12   Figure 1.2: Structure and cross-linking of RG-II. (A) RG-II contains polygalacturonan backbone and four distinct side-chains (A to D). (B) Schematic of RG-II cross-linking into a dimer through borate. (C) Apioses from A side-chain cross-linked through borate diester linkage. Reprinted with permission under CC-BY 4.0 licence (https://creativecommons.org/licenses/by/4.0/). Bar-Peled, M., Urbanowicz, B.R., and O’Neill, M.A. (2012). The Synthesis and Origin of the Pectic Polysaccharide Rhamnogalacturonan II – Insights from Nucleotide Sugar Formation and Diversity. Frontiers in Plant Science 3: 1–12. https://www.frontiersin.org.  13   1.1.4 Hydroxyproline-rich glycoproteins Aside from enzymes, cell walls also contain proteins that perform structural roles. Structural proteins of plant cell walls are usually grouped by their amino-acid composition into hydroxyproline-rich glycoproteins (HRGPs), proline-rich proteins (PRPs) and glycine-rich proteins (GRPs) (Josè and Puigdomènech, 1993). However, HRGPs are the only group with defined roles in plant cell walls. These proteins are highly glycosylated with arabinan/arabinogalactan glycans similar in composition to RG-I side-chains (Nguema-Ona et al., 2014). In comparison to wide-spread N-glycosylation of asparagines (Asn) in secreted proteins, these proteins are characterised by O-glycosylation at hydroxyproline (Hyp) or, less frequently, serine (Ser) amino acids in the peptide (Nguema-Ona et al., 2014). Hyp residues are initially proline (Pro) residues that become hydroxylated in the ER and Golgi apparatus, followed by attachment of the glycans (Nguema-Ona et al., 2014). Two of the major HRGP groups are extensins (EXTs) and arabinogalactan proteins (AGPs) distinguished by differences in glycan composition, glycan content, amino acid motifs, and biological function.    14   Figure 1.3: Schematic representation of typical O-glycosylation patters of plant HRGPs. (A) Type II arabinogalactan glycan attached to hydroxyproline of an AGP. (B) Typical glycosylation pattern of Ser-Hyp-Hyp-Hyp-Hyp motif from EXTs. Reprinted with permission under CC-BY 4.0 licence (https://creativecommons.org/licenses/by/4.0/). Nguema-Ona, E., Vicré-Gibouin, M., Gotté, M., Plancot, B., Lerouge, P., Bardor, M., and Driouich, A. (2014). Cell wall O-glycoproteins and N-glycoproteins: aspects of biosynthesis and function. Frontiers in Plant Science 5: 499. Panel A of Figure 1 from the publication was cropped out, and panels B and C re-labelled as A and B. https://www.frontiersin.org.  1.1.4.1 Extensins Extensins are HRGPs with Ser-Pro-Pro-Pro-Pro amino acid repeats, where Pro residues get hydroxylated to give a Ser-Hyp-Hyp-Hyp-Hyp motif, followed by O-glycosylation of these residues (Figure 1.3 B; Kieliszewski and Lamport, 1994; Nguema-Ona et al., 2014). The glycosylation involves the attachment of a single α-D-Gal residue on Ser (Saito et al., 2014), and 15  β-L-Araf , either as a single residue or short (1→2) and (1→3)-linked arabinan oligosaccharides, on contiguous Hyp residues (Figure 1.3 B; Velasquez et al., 2011; Saito et al., 2014). EXTs are important for cell wall assembly and morphogenesis, as suggested by developmental and cell shape defects in mutants with aberrant EXTs (Cannon et al., 2008; Velasquez et al., 2011). These proteins are known to be cross-linked through tyrosines (Tyr) by peroxidases to create covalent cross-links that promote cell wall strengthening (Waffenschmidt et al., 1993; Cannon et al., 2008), but defects in mutants affecting glycosylation of EXTs suggest that glycan chains are also necessary for their function (Velasquez et al., 2011). It has been proposed that glycans maintain EXTs in an extended form necessary for cross-linking (Stafstrom and Staehelin, 1986). The cross-linking properties of EXTs are used as a response to pathogen attack to promote resistance, further demonstrating the importance of these proteins (Deepak et al., 2010).  1.1.4.2 Arabinogalactan proteins O-glycosylation in AGPs occurs on Hyp residues that are either contiguous or non-contiguous (Kieliszewski et al., 2011). AGPs have glycans much larger in size than EXTs, which contain Gal mixed with Ara into type II arabinogalactans (Figure 1.3 A). The backbone of these chains is linear (1→3)-β-D-Gal, with Gal branching at the C-6 into short β-(1→6)-galactans, which in turn branch into single α-L-Araf residues or short α-(1→5)-Araf chains at C-3 (Figure 1.3 A; Tryfona et al., 2012). Terminal L-Fucp and L-Rhap residues were also detected in AGP glycans, as well as D-GlcpUA, which, similar to glucuronoxylans, is often methoxylated at the C-4 (Tryfona et al., 2012; Nguema-Ona et al., 2014). Most AGPs are membrane-anchored through glycosylphophatidylinositol (GPI) anchors at the C-terminus of the protein, which are often cleaved (Schultz et al., 2004; Ellis et al., 2010). The exact roles of AGPs are less well understood 16  than those of EXTs, but they have been proposed to influence cellulose biosynthesis (MacMillan et al., 2010), development of both female (Acosta-García and Vielle-Calzada, 2004) and male gametophyte (Coimbra et al., 2009), fertilisation (Wu et al., 1995; Cheung et al., 1995),  and signalling (Xu et al., 2008; Van Hengel and Roberts, 2003; Motose et al., 2004).  1.1.5 Callose Callose is a linear β-(1→3)-glucan (Figure 1.1), sometimes with β-(1→6) branches, which often appears in cell walls transiently (Chen and Kim, 2009; De Storme and Geelen, 2014). It first appears in high abundance during the cytokinesis at the time of cell plate formation. It has been proposed that callose assists the membrane in the cell plate expansion, after which it is degraded by β-(1→3)-glucanases (Samuels et al., 1995; Thiele et al., 2009). This polysaccharide is also involved in the regulation of the size exclusion limit of plasmodesmata and symplastic transport of molecules from cell to cell (De Storme and Geelen, 2014). It is synthesised within the neck of plasmodesmata to increase the size exclusion, whereas its absence can reduce the size exclusion limit, allowing larger molecules, such as transcription factors, mRNAs, or viruses, to pass through (Guseman et al., 2010; Vatén et al., 2011; Li et al., 2012). Another role of great importance is the regulation of phloem transport, as suggested by problems with sugar transport of mutants lacking the callose lining around the pores of sieve elements (Barratt et al., 2011). Callose appears in the pollen wall during its development, and mutants defective in callose deposition during this process have collapsed, non-viable, pollen grains (Töller et al., 2008). It has been proposed to prevent fusion of microspores after meiosis and ensure their separation into individual developing pollen grains (Dong et al., 2005; Shi et al., 2016). Pollen tubes also deposit 17  callose between the original pollen grain and the sperm cells, but this deposition does not seem to be essential for fertilisation (Parre and Geitmann, 2005; Nishikawa et al., 2005).  1.1.6 Primary cell wall assembly, structure, and organisation Primary cell wall assembly in eudicots and monocots, other than Poales, starts in cell division during cell plate formation, when pectin, xyloglucan and callose are deposited (Verma, 2001; Northcote et al., 1989; Moore and Staehelin, 1988; Baluška et al., 2005). Matrix polysaccharides, pectins and XyG, are synthesised in the Golgi apparatus (Rybak et al., 2014) or endocytosed from the existing cell wall (Baluška et al., 2005; Dhonukshe et al., 2006), and trafficked inside vesicles to the growing cell plate. In contrast, callose is synthesised at the membrane directly into the growing cell plate (Drakakaki, 2015). HG is synthesised mostly in the methyl-esterified form, and later gets de-esterified in the apoplast by pectin methylesterases (Rybak et al., 2014). Cellulose starts being synthesised within the matrix as the cell plate matures, replacing callose that is degraded (Drakakaki, 2015; Samuels et al., 1995). The cell wall matrix keeps cellulose microfibrils apart and regulates the degree of their aggregation/self-association (McCann et al., 1990; Anderson et al., 2010). Hemicelluloses interact with the surface of the microfibrils through hydrogen bonding (Valent and Albersheim, 1974). This is promoted by the conformational similarities between cellulose and hemicelluloses, where linear hemicellulosic polymers often resemble cellulose and tend to associate with it, and branching reduces those associations (Kabel et al., 2007; Köhnke et al., 2011; Busse-Wicher et al., 2016). On the other hand, pectic polymers can interact with cellulose through arabinan and galactan side-chains of RG-I (Zykwinska et al., 2005). Similar to hemicelluloses, fewer side-chains are correlated with stronger associations with microfibrils (Zykwinska et al., 2005). Such interactions, however, appear to be weaker than those 18  between XyG and cellulose (Zykwinska et al., 2005), most likely due to the fact that the side-chains are shorter and their conformation results in less efficient alignment with cellulose (Zykwinska et al., 2006). RG-I and hemicelluloses (XyG and xylans) can be covalently linked (Duan et al., 2010, 2004; Popper and Fry, 2008; Tan et al., 2013), but it is not known whether these cross-links are made during their synthesis or following secretion to the apoplast. The ability to extract pectin with water, chelators or Na2CO3 suggests that only a fraction is covalently linked to hemicelluloses, since extraction of hemicelluloses requires strong bases.  Hemicelluloses, namely XyG, can be re-modelled by xyloglucan endotransglucosylases (XETs). These enzymes break the glucan backbone of XyG and create new glycosidic bonds to another XyG molecule (Thompson and Fry, 2001). As such, newly secreted XyG can get integrated into an already existing network. Extensins have been proposed to be involved in the assembly of a new cell wall, inferred from mutant phenotypes (Cannon et al., 2008). Based on 2D-NMR studies of the cell wall, the protein network appears to be separate from the carbohydrate network (Wang et al., 2012), but direct covalent linkages between the glycans of HRGPs and matrix polysaccharides have also been shown (Tan et al., 2013). EXTs are thought to be cross-linked through Tyr residues by the action of peroxidases and H2O2, creating Tyr dimers, trimers or tetramers (Fry, 2004). Dimers can be dityrosines, where phenyl rings of two tyrosines are linked by a C-C bond (biphenyl linkage), or isodityrosines (IDTs), where they are linked through an ether bond (diphenyl ether linkage; Figure 1.4 (a); Fry, 1982). However, dityrosines have never been isolated from plants, suggesting that IDTs may be the only form existing in vivo. IDTs occur both as intramolecular and intermolecular cross-links (Fry, 1982; Brady and Fry, 1997; Held et al., 2004). Pulcherosine 19  has been isolated as a trimer of tyrosines, containing both biphenyl and diphenyl ether linkages (Figure 1.4 (b); Brady et al., 1998). A tetramer present in the cell walls is di-isodityrosine (di-IDT), a dimer of IDTs linked through a C-C bond (Figure 1.4 (c); Brady et al., 1996). It is, therefore, possible that IDT cross-links to a single Tyr to form a pulcherosine cross-link, or it can cross-link to another IDT to form di-IDT cross-link (Figure 1.4; Brady and Fry, 1997). It is also possible that pulcherosine cross-links to Tyr to form di-IDT (Figure 1.4). Aside from cross-linking through IDTs, peroxidases can cross-link EXTs through different amino acids. Tyr-Lys cross-link has been suggested as another way of cross-linking (Schnabelrauch et al., 1996).  Figure 1.4: Oxidative cross-linking of HRGPs through tyrosines (Tyr). Figure based on Fry, 2004.  Another type of oxidative cross-linking occurs between hydroxycinnamic acids, mainly ferulic acid (FA). Hydroxycinnamic acids were found to be ester-linked to Gal or Ara in side-chains of RG-I (Saulnier and Thibault, 1999; Ralet et al., 2005), or to Ara side-chains of grass arabinoxylans (Saulnier and Thibault, 1999; Encina and Fry, 2005; Grabber et al., 1995). Hydroxycinnamates can be released from the cell wall through saponification of ester bonds by bases, which allow isolation and analysis of the dimers naturally occurring in the cell walls. The most common dimers of FA are C-C-linked through C-8 and C-5 positions (8-5’ diferulates; 20  Figure 1.5), 8-8’ (Figure 1.5), and 5-5’ (Figure 1.5) diferulates. Cross-linking can also occur through a hydroxy group to create ether linkages, with the most common being 8-O-4’ (Figure 1.5), and the most rare being 4-O-5’ (Figure 1.5; Bunzel et al., 2001; Saulnier and Thibault, 1999; Ralph et al., 1994). FA trimers have also been isolated from the cell walls of maize bran (Figure 1.5, structure 10; Bunzel et al., 2006). Besides FA, other hydroxycinnamic acid dimers have been isolated from the cell walls, namely sinapate ester dimers and mixed ferulate-sinapate ester dimers (Bunzel et al., 2003). Two isolated disinapates are cyclic (Figure 1.5) and noncyclic (Figure 1.5) 8-8’ dimers, whereas sinapate-ferulate heterodimers were found in 8-8’ cyclic, 8-8’ noncyclic, 8-5’ noncyclic, and 8-O-4 forms (Bunzel et al., 2003). Even though p-coumarate esters are present in grass cell walls, their dimers have not been detected and their roles are not known (Hatfield et al., 2017). One mechanism of wall-bound hydroxycinnamate cross-linking in primary cell walls involves peroxidases and H2O2. The polysaccharides are feruloylated intracellularly (Obel et al., 2003; Fry, 1987) by an unknown mechanism (Hatfield et al., 2017) and cross-linked by peroxidases in the cell wall (Encina and Fry, 2005). It is unclear whether these cross-links contribute to structural support or cell adhesion, but they appear to contribute to resistance against cell wall degradation by fungal enzymes (Grabber et al., 1998). 21   Figure 1.5: Structures of dimers and trimers of hydroxycinnamic acids isolated from plant cell walls. Figure based on Bunzel et al., 2004.  22  1.2 Arabidopsis seed coat mucilage Seed coat mucilage is an extracellular matrix rich in pectin, which contains all the structural components of type I cell walls: cellulose, hemicelluloses, pectins, and structural proteins. It is synthesised by seed coat epidermal cells and deposited to the apoplast beneath the primary cell wall at the seed surface. When the seed is exposed to water it swells, breaks the primary wall and extrudes to form a capsule around the seed (Figure 1.6 A). Mucilage has emerged as a model to study cell walls due to three main reasons (Haughn and Western, 2012). First, unlike other cell types where defects in cell walls often have severe developmental and/or physiological abnormalities, seeds that completely lack mucilage are relatively unaffected. This makes mucilage a good candidate for genetic manipulations. Second, mucilage is highly abundant and easily extractable, which makes it easy to analyse its polysaccharide composition. Third, for research purposes, cell wall material is extracted from tissues containing multiple cell types making it hard to determine which cell wall components are derived from which cell type.   Figure 1.6: Hydrated wild-type seeds (Col-2) of Arabidopsis thaliana stained with Ruthenium Red. (A) Seed stained directly with Ruthenium Red. Both non-adherent mucilage and adherent mucilage surround the seed. (B) Seed stained after shaking in water. Only adherent layer remains around the seed. Bars = 200 μm. 23  1.2.1 Mucilage is produced from a single cell type Seed coat development begins after fertilisation (Haughn and Chaudhury, 2005). Signals from the endosperm stimulate differentiation of seed coat cells from cells of the ovule integuments (Garcia et al., 2003). The outermost layer of ovule integuments undergoes differentiation into the seed coat epidermis, involving mucilage deposition and development of a thick cellulosic secondary cell wall called the columella (Western et al., 2000; Windsor et al., 2000). In Arabidopsis, anthesis is a process where anthers with mature pollen grow to the height of the stigma of the pistil to allow for pollination. This occurs immediately before fertilisation. For this reason, the progression of seed coat development is estimated using days post anthesis. Therefore, 0 days post anthesis (DPA) represents the time at which the anthesis, and pollination, occurred. Following double fertilisation, the seed coat epidermal cells start elongating rapidly to accommodate later growth of the embryo and endosperm (Western et al., 2000). At these stages (0-4 DPA), much of the volume of the epidermal cell is occupied by a large vacuole (Figure 1.7 A to B). Once growth has ceased (4-5 DPA; Figure 1.7 B), the cell accumulates starch granules inside amyloplasts in the cytosol until 7 DPA (Figure 1.7 B to D). At 5 DPA, mucilage deposition at the junction of the radial and outer wall forming is initiated forming a doughnut shaped mucilage pocket (Figure 1.7 C). From 5 DPA to 9 DPA, continuing mucilage deposition results in the expansion of the doughnut-shaped mucilage pocket (Figure 1.7 D). Concomitantly, the vacuole contracts toward the bottom of the cell leaving a volcano-shaped cytoplasmic column in the centre.  Following the completion of mucilage deposition (9 DPA), the synthesis of the columella begins with deposition of cellulose outside the membrane of the cytoplasmic column, displacing the cytoplasm and membrane toward the bottom of the cell (Figure 1.7 E). Throughout differentiation the amyloplasts decrease in size and number, suggesting that the 24  glucose for polysaccharide synthesis comes from the starch in the amyloplasts. By the end of the columella synthesis (13 DPA) the cytoplasm disappears resulting in a cell comprised of a primary wall and a columella with mucilage between the two (Figure 1.7 F). Finally, the cell undergoes programmed cell death and mucilage dehydrates. Once the mature seed is exposed to water, the mucilage hydrates and expands (Figure 1.7 G), rupturing the primary cell wall (Figure 1.7 H).   Figure 1.7: Developmental sequence of a seed coat epidermal cell. Schematics represent a side view (cross-section) of a single cell. Figure based on North et al., 2014.  1.2.2 Composition of the seed coat mucilage When dry, mature, seeds are hydrated, they rapidly extrude mucilage that forms a halo around the seed. The extruded mucilage forms two distinct layers: an outer, non-adherent layer, and inner, adherent layer (Figure 1.6 A; Haughn and Western, 2012). The non-adherent layer is easily removed by gentle shaking (Figure 1.6 B), whereas the adherent layer requires extensive vigorous shaking, sonication, enzymes or strong bases to be removed, suggesting compositional and/or structural differences between the two. The main component of both layers is RG-I (Macquet et al., 2007a; Dean et al., 2007). The biosynthesis of RG-I backbone requires RG-I:RHAMNOSYLTRANSFERASE1 (RRT1), which adds rhamnose into the growing 25  polysaccharide (Takenaka et al., 2018). Mucilage RG-I appears to be synthesised with β-(1→5)-arabinans and mostly single Gal as its side-chains (Dean et al., 2007; Arsovski et al., 2009). Gal side-chains are removed by a β-galactosidase MUCILAGE-MODIFIED2 (MUM2; Dean et al., 2007; Macquet et al., 2007b), whereas arabinans are removed by a bi-functional β-D-xylosidase/α-L-arabinofuranosidase BETA-XYLOSIDASE1 (BXL1; Arsovski et al., 2009) following secretion to the apoplast. These modifications result in mostly un-branched RG-I (Figure 1.8). The non-adherent mucilage layer is composed almost exclusively of RG-I, whereas the adherent layer also contains cellulose, xylans, galactoglucomannans, HG, and structural proteins (Voiniciuc et al., 2015c; Tsai et al., 2017). Cellulose is synthesised by CESA complexes containing subunits commonly associated with primary cell wall formation: CESA3, CESA5 and most likely CESA1 (Griffiths et al., 2015; Mendu et al., 2011). Starting at 6-7 DPA, these CESA complexes move around the cytoplasmic column in a unidirectional manner, depositing cellulose (Griffiths et al., 2015).  The hemicelluloses found in mucilage are xylans with xylose branches (Figure 1.8), and galactoglucomannans (Figure 1.8). IRREGULAR XYLEM7 (IRX7) is involved in the biosynthesis of the xylan through an unknown mechanism (Hu et al., 2016b), whereas the xylan backbone is synthesised by IRREGULAR XYLEM14 (Figure 1.8; IRX14; Hu et al., 2016a; Voiniciuc et al., 2015a), and side-chain xyloses are added onto the backbone by MUCILAGE-MODIFIED5/MUCILAGE-RELATED21 (Figure 1.8; MUM5/MUCI21; Voiniciuc et al., 2015a). Galactoglucomannan requires CELLULOSE SYNTHASE-LIKE A2 (CSLA2; Yu et al., 2014) for biosynthesis of the backbone with alternating Glc and Man residues (Yu et al., 2018), and MUCILAGE-RELATED10 (MUCI10; Yu et al., 2018; Voiniciuc et al., 2015b) for addition of the Gal side-chains (Figure 1.8). In total, HG makes about 10% of the mucilage (Voiniciuc et al., 2015c). The degree of HG methylesterification (DM) in the mucilage 26  is close to 35% (Yu et al., 2014). Based on immunolabelling experiments, lower DM HG is present in the adherent mucilage, whereas higher DM HG is distributed at the edges of the adherent mucilage (Rautengarten et al., 2008; Macquet et al., 2007a). Cell wall proteins were detected in both layers of the mucilage (Tsai et al., 2017), but none of the mutants in genes encoding those proteins had visible mucilage phenotypes. The only putative structural protein with a defined biological role, SALT-OVERLY SENSITIVE5 (SOS5; Harpaz-Saad et al., 2011) was not detected in the mucilage proteome. As suggested by the genetic analysis, SOS5 is a fasciclin-like AGP (FLA) with roles in mucilage adherence independent of CESA5 (Griffiths et al., 2014).  Figure 1.8: Polysaccharide components of the Arabidopsis seed coat mucilage. Arrows indicate monosaccharides and glycosyltransferases responsible for their addition to the polysaccharide chains.  27  1.2.3 Structure of the seed coat mucilage The adherent layer of mucilage has a very distinct structure. Even though it is mostly composed of RG-I, its structure is defined by cellulose. Cellulose appears in two forms, as cellulosic rays and as diffuse cellulose (Griffiths et al., 2014). A distinct cellulosic ray appears attached to the top of the columella after mucilage extrusion. Circular unidirectional movement of CESA5 and CESA3 around the cytoplasmic column suggests that cellulose is deposited into the mucilage in linear arrays around the columella (Griffiths et al., 2015). Upon extrusion the cellulose unwinds to form a ray. Rays contain mucilage hemicelluloses, xylans and GGMs that surround and bind to the cellulose (Yu et al., 2014; Voiniciuc et al., 2015b; Ralet et al., 2016; Voiniciuc et al., 2015a; Hu et al., 2016a; Yu et al., 2018).  Immunolabelling experiments indicate that partially methylesterified HG is also present in the rays (Griffiths et al., 2014). Therefore, cellulosic rays appear to contain all the components of the adherent mucilage organised around cellulose microfibrils. Diffuse cellulose, as the name suggests, is less organised, and appears in spaces between the rays (Griffiths et al., 2014). Based on these two types of cellulose organisation, adherent mucilage can be separated into cellulosic rays and the space between the rays.  1.2.4 Function of cell wall components in the mucilage CESA5 is a key subunit of the CESA complex that synthesises mucilage cellulose. Mutations in CESA5 results in a drastic decrease in the amount of mucilage cellulose and the loss of adherence (Mendu et al., 2011; Harpaz-Saad et al., 2011; Sullivan et al., 2011), suggesting that the cellulose scaffold has a role in anchoring the mucilage to the seed surface.    Like those of CESA5, mutants affecting mucilage xylans, irx7, irx14 and mum5/muci21, also exhibit loss of mucilage adherence (Hu et al., 2016b, 2016a; Voiniciuc et al., 2015a; Ralet et 28  al., 2016). Xylans interact with cellulose and are responsible for mediating the adherence of mucilage RG-I (Ralet et al., 2016). Since covalent bonds between xylans and RG-I have been described (Tan et al., 2013) it is possible that mucilage adherence is mediated by a xylan-RG-I polysaccharide that binds to mucilage cellulose.  Mutations in several other genes also impact mucilage adherence although their role in adherence is not well understood. SOS5 appears to influence adherence through pectin rather than cellulose directly (Griffiths et al., 2014). It is secreted to the mucilage pocket during seed coat epidermal cell differentiation (Griffiths et al., 2016), but it is not detected in proteomic analysis of mature mucilage (Tsai et al., 2017), suggesting that it is not a structural component. Receptor-like kinase FEI2 appears to be functioning in the same pathway as SOS5 (Griffiths et al., 2016; Harpaz-Saad et al., 2011), but little is known about the mechanism in which they function. Thus, how SOS5 influences mucilage structure and function remains an open question.   GGMs are needed for normal ray structure. Mutations that alter or reduce GGM result in more compact mucilage with shorter rays (Voiniciuc et al., 2015b; Yu et al., 2014) indicating a role in establishing the structure and scaffolding of the mucilage. The binding of GGM to the cellulose ray microfibrils may reinforce them. Alternatively, the GGMs may coat the microfibrils and limit hydrogen bonding between adjacent microfibrils within the ray to maintain its structure.  Hints to the role of HG arise from the phenotypes observed in mutants with changes in DM of HG in mucilage. The low DM in flying saucer1 (fly1) mutant is associated with detachment of the primary cell wall from the columella and a more compact adherent mucilage capsule, whereas reduced DM in primary cell walls of sbt1.7/ara12 (a mutant for gene encoding a protease) and pectin methylesterase inhibitor6 (pmei6; a mutant for gene encoding a PME 29  inhibitor) is correlated with the failure to extrude mucilage properly (Rautengarten et al., 2008; Voiniciuc et al., 2013; Saez-Aguayo et al., 2013).  Since mucilage does not appear to be less adherent in these mutants, Ca2+ bridges between HG molecules seem to be participating in mediating mucilage coherence, which was demonstrated by shrunken halo of CaCl2-treated fly1 seeds (Voiniciuc et al., 2013).  1.2.5 Removal of RG-I side-chains is needed for extrusion One intriguing question that remains to be answered is how RG-I side-chains and their modifications affect its properties. MUM2 and BXL1, genes that encode enzymes responsible for the removal of RG-I side-chains following its secretion to the mucilage pocket. The removal of the side-chains is essential for normal mucilage expansion and extrusion when seeds are exposed to water, since the mucilage of mum2 and bxl1 mutants cannot expand properly and does not extrude normally (Figure 1.9; Arsovski et al., 2009; Macquet et al., 2007b; Dean et al., 2007). An explanation for how the side-chains influence mucilage expansion has eluded researchers for over a decade.  30   Figure 1.9: Phenotypes of mucilage-modified2 (mum2). (A) and (B) Seeds hydrated in water and stained with Ruthenium Red. (A) Wild type (Col-2); (B) mum2-1; (C) and (D) Schematics showing differences between RG-I structure in wild type (C) and mum2 (D) mucilage based on Macquet et al., 2007b and Dean et al., 2007. Bars = 200 μm.  1.3  Research questions and goals It is evident that MUM2 strongly affects RG-I hydration properties, but the underlying mechanism is not understood. It is possible that the addition of Gal to RG-I affects its intrinsic hydration properties, but the addition of Gal to mannans has the opposite effect (increased hydration properties). To address this question, we undertook a genetic suppressor approach to identify genes influencing mucilage expansion. One mutant was isolated from this screen, named ruby particles in mucilage (ruby) fully suppressed the mum2 extrusion phenotype. RUBY was 31  mapped to At1g19900, a gene annotated to encode a glyoxal oxidase-related enzyme. RUBY is a member of a small gene family, but none of the family members have been characterized.  My doctoral thesis research had two main research objectives: 1. To determine how RUBY acts to influence mucilage expansion. 2. To investigate the roles of RUBY homologues.  32  Chapter 2: Materials and Methods 2.1 Plant materials and growth conditions 2.1.1 Arabidopsis thaliana Arabidopsis thaliana plants were grown in growth rooms or growth chambers under continuous light at 20-22°C and low light intensity (80-120 μmol m-2 s-1). Seeds were surface-sterilised with 70% EtOH and germinated on AT minimal medium (Haughn and Somerville, 1986) with 0.7% w/v agar. Seedlings were transferred to soil mix (Sunshine Mix #4, Sun Gro Horticulture) once true leaves formed. Table 2.1: A list of mutant lines used in this study. Gene Line Allele Mutation type Ecotype Publication RUBY  ruby-1 EMS Col-2 This study RUBY  ruby-2 EMS Col-2 This study RUBY  ruby-3 EMS Col-2 This study RUBY SALK_020627 ruby-4 T-DNA insertion Col-0 Alonso et al. (2003) RUBY WiscDsLoxHs097_11H ruby-5 T-DNA insertion Col-0 Woody et al. (2007) MUM2  mum2-1 EMS Col-2 Western et al. (2001); Dean et al. (2007) MUM2 SALK_011436 mum2-10 T-DNA insertion Col-0 Alonso et al. (2003); Dean et al., (2007) BXL1 CS16299 bxl1-1 T-DNA insertion Ws-2 Arsovski et al. (2009)   33  Table 2.1: A list of mutant lines used in this study. AOMT1 GK-007F02-014809 aomt1-1 T-DNA insertion Col-0 Kleinboelting et al. (2012); Fellenberg et al. (2012) COMT1 SALK_002373 comt1-1 T-DNA insertion Col-0 Vanholme et al. 2012 4CL1 SAIL_350_H10 4cl1-3 T-DNA insertion Col-3 Sessions et al. (2002) 4CL3 SALK_003025 4cl3-3 T-DNA insertion Col-0 Alonso et al. (2003) F5H  fah1-2 EMS Col-0 Chapple et al. (1992) F5H  fah1-7 EMS Ler  REF1  ref1-4 EMS Col-0 Ruegger and Chapple (2001) GOXL1 SALK_203669 goxl1-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL1 SALK_000947 goxl1-2 T-DNA insertion Col-0 Alonso et al. (2003) GOXL1 SALK_000955 goxl1-3 T-DNA insertion Col-0 Alonso et al. (2003) GOXL1  goxl1-4 1 bp deletion Col-2 This study GOXL2 SALK_142573 goxl2-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL3 SALK_046326 goxl3-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL4 SALK_093271 goxl4-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL4 SAIL_33_A07 goxl4-2 T-DNA insertion Col-3 Sessions et al. (2002)  34  Table 2.1: A list of mutant lines used in this study. GOXL4 SM_3_34763 goxl4-3 transposon insertion Col-0  GOXL5 SALK_042652 goxl5-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL5 SAIL_310_B02 goxl5-2 T-DNA insertion Col-3 Sessions et al. (2002) GOXL6 SALK_022199 goxl6-1 T-DNA insertion Col-0 Alonso et al. (2003) GOXL6 SAIL_840_G10 goxl6-2 T-DNA insertion Col-3 Sessions et al. (2002) GOXL6 GT_3_5998 goxl6-3 transposon insertion Ler  GOXL6 SAIL_314_C08 goxl6-4 T-DNA insertion Col-3 Sessions et al. (2002) GOXL6  goxl6-5 2 bp insertion Col-2 This study  ruby-1, ruby-2 and ruby-3 were isolated from an EMS-mutagenised mum2-1 mutant population, which was generated in Col-2 ecotype background (Western et al., 2001). ruby-4, ruby-5, aomt1-1, comt1-1, 4cl1-3, 4cl3-3, fah1-2,  fah1-7, goxl1-1, goxl1-2, goxl1-3, goxl2-1, goxl3-1, goxl4-1, goxl4-2, goxl4-3, goxl4-4, goxl5-1, goxl5-2, goxl6-1, goxl6-2, goxl6-3 and goxl6-4 were ordered from Arabidopsis Biological Resource Center (ABRC) through TAIR (www.arabidopsis.org). ref1-4 was a gift from Dr. Clint Chapple (University of Purdue).  2.2 Statistical Analyses In the analyses, a biological replicate represents a batch of seeds harvested from 6 plants of the same genotype from the same pot, grown in the same tray with other genotypes employed in the analysis. Separate biological replicates were grown under the same conditions at different times to ensure statistical independence and account for variations between separate growth trials. 35  Statistical tests were done in R (https://www.r-project.org/) using RStudio (https://www.rstudio.com/). Tukey’s HSD test was done after Welch’s One-way or Two-way ANOVA, and the letters designating groups were generated using “multcompView” package. Pair-wise comparisons were done using Welch’s t-test. Data frames for plotting were arranged using “plyr” and “reshape2” packages, and results were plotted using “ggplot2” package.  2.3 Microscopy 2.3.1 Light microscopy For light microscopy of mucilage, dry seeds were shaken in water for 2 h, washed twice to remove non-adherent mucilage, stained with 0.02% (w/v) Ruthenium Red (Cat# R2751, Sigma) for 10 min, and washed twice in water. They were imaged using Leica DFC450 C camera (Leica Microsystems) attached to Zeiss AxioSkop2 light microscope (Carl Zeiss). Imaging of the primary cell wall detachment was done with seeds shaken in water for 1 h using the same microscope with DIC configuration. Filming of mucilage extrusion was completed on a single seed placed on a microscope slide. It was covered with a coverslip, which was held in place by the weight of metal forceps to prevent movement of the seed during filming. 0.02% (w/v) Ruthenium Red was added under the coverslip immediately after the start of filming. ProGOXL:GUS plants were imaged using Carl Zeiss™ Stemi 2000-C stereo microscope (Carl Zeiss) with Canon EOS Rebel T5 camera (Canon Canada Inc.) or Nikon SMZ18 stereo microscope (Nikon Instruments Inc.) for larger organs, and Zeiss AxioSkop2 light microscope (Carl Zeiss) for smaller organs and sections. Siliques of plants with goxl1-4 and goxl6-5 36  mutations were de-stained in EtOH/AcOH 1:1 (v/v), and imaged using Nikon SMZ18 stereo microscope (Nikon Instruments Inc.).  2.3.2 Spinning disk confocal microscopy Cellulose on the seed surface was stained using Calcofluor White M2R (CFW; Cat# F3543, Sigma) or Pontamine Fast Scarlet 4B (Cat# S479896, Sigma), 0.1 M NaCl. Seeds were shaken for 1h in water, washed twice, shaken in CFW/Pontamine for 1 h protected from light, washed twice in water and kept in the dark until imaged. Seeds were imaged using Hamamatsu C9100-02 CCD camera (Hamamatsu Photonics) attached to a Leica DMI6000 inverted microscope (Leica Microsystems) combined with PerkinElmer UltraVIEW VoX Spinning Disk Confocal system (PerkinElmer), with excitation at 405 nm and 460/50 nm emission filter for CFW, or excitation at 561 nm, emission filter 595/50 nm for Pontamine S4B. For imaging of expression and localisation in developing seeds, the flower most recently opened [corresponds to 0 days post anthesis (DPA)], on T2 and T3 plants expressing Citrine fusion constructs, were marked using washable water-soluble paint. Valves of siliques at the desired developmental stage were peeled using forceps and seeds mounted on a glass slide with water. Seeds were imaged using the same spinning-disk confocal system as for CFW staining, except that excitation and emission were set at 514 nm and 540/30 nm, respectively. For plasma membrane co-localisation, seeds of the same plants were first incubated in 10 μM FM4-64 (Cat# T-3166, Thermo Fisher), vacuum-infiltrated for 5 min, incubated in the dark for another 10 min, and washed in water. After mounting on slides, they were imaged at excitation 514 nm, emission filter 540/30 nm for Citrine, and 561 nm excitation laser and 650/75 nm emission filter for FM4-64. 37  For exine staining, mature pollen grains were stained in 0.001% (w/v) Auramine O (Cat# 861030, MilliporeSigma) in 50 mM Tris-HCl buffer pH 7.5 as described by Dobritsa et al., 2009, covered with a coverslip, sealed using nail polish and imaged at 488 nm excitation, 525/36 nm emission using the spinning disc confocal microscope described above.  2.3.3 Scanning electron microscopy (SEM) Seed surface was imaged with Hitachi S4700 scanning electron microscope (Hitachi High-Technologies), after coating dry seeds with Au/Pd using EMPrep2 sputter coater (Nanotech).  2.4 Genotyping Arabidopsis leaf DNA was collected by pressing leaves onto FTA cards (Whatman). For genotyping, small FTA card discs were washed in FTA buffer (10 mM Tris-HCl pH 7.5, 2 mM EDTA, 0.1% [v/v] Tween 20) followed by two washes in TE-1 buffer (10 mM Tris-HCl pH 8, 0.1 mM EDTA). Polymerase chain reactions (PCRs) were performed on FTA discs using primers listed in Table 2.2. Table 2.2: A list of primers used to genotype mutations used in this study. Primer Name 5ʹ-3ʹ Sequence Allele Additional Information MUM2_87f GAAACAATGCGTATAGCTTGAG mum2-1 TatI restriction digestion MUM2_77r ACAGGGTTCTAGTATCAGTAAACGTC MUM2_88f GACAACTCTAACACAATCACG mum2-10  MUM2_79r GCATTTCATCCCGTTGCAGG LBb1.3 ATTTTGCCGATTTCGGAAC GOXL_55f AACGATGCCAAACCTGAATGGG ruby-1 MseI restriction digestion GOXL_56r CGTCCCATATTTAATCATCGACTG 38  Table 2.2: A list of primers used to genotype mutations used in this study. GOXL_13f TCCAAAATCGACTGCACgGC ruby-2 Wild-type allele GOXL_19r CAAAAACGGTAACGCGACTAC GOXL_20f TCCAAAATCGACTGCACgGt Mutant allele GOXL_19r CAAAAACGGTAACGCGACTAC GOXL_1f GACACAAGTCACAGTACATTCTC ruby-3 BstBI restriction digestion GOXL_19r CAAAAACGGTAACGCGACTAC SALK_020627-RP ATCAGAGGGTTTTGGTTTTGG ruby-4  GOXL_4r CTTAATACAGCAAATTATCAATGATC LBb1.3 ATTTTGCCGATTTCGGAAC ruby-5f ACAGTTCTGTAATGTTAGGAATCG ruby-5  GOXL_51r CCGTTAAAGGTCGGATGGTG WDLH_L4new GTAGATTTCCCGGACATGAAG bxl1-1_3f GTCTATCAAATAAGTCCGAAACG bxl1-1  bxl1-1_4r TGAAGTCCCCGGACGTAGC Feldman_LB GATGCACTCGAAATCAGCCAATTTTAGAC comt1-1LP ATGCCTCAAACTCTTTCTCGG comt1-1  comt1-1RP TGACTTCTTTGGTTGATGTTGG LBb1.3 ATTTTGCCGATTTCGGAAC aomt1-1LP AAAGCAAAACAAGGACAATGG aomt1-1  aomt1-1RP GCTCACAAGATCGACTTCAGG GK-8409 ATATTGACCATCATACTCATTGC 4cl1-3LP GTTTTGCCCTCAGATCTTTCC 4cl1-3  4cl1-3RP AAGGTTACCTCAACAATCCGG LB3 TAGCATCTGAATTTCATAACCAATCTCGATACAC 4cl3-3LP TTTTGGCAAGTACTAATTCGC 4cl3-3  4cl3-3RP TCGCAACTACAAAGGATACTGC LBb1.3 ATTTTGCCGATTTCGGAAC   39  Table 2.2: A list of primers used to genotype mutations used in this study. goxl1-1 LP2 TTGGATTTGATGATTATCGCC goxl1-1  goxl1-1 RP3 AAACAAAGTGCTCAAAGAGTTCCC LBb1.3 ATTTTGCCGATTTCGGAAC goxl1-2 LP ATTTAACGCTGTAAGCCAACG goxl1-2  goxl1-2 RP ACCGTTTACGGTTAAACCTCC LBb1.3 ATTTTGCCGATTTCGGAAC goxl1-3 LP CCAGGCAAAATTTCAAAAGTG   goxl1-3  goxl1-3 RP CCTCCAGAAGAACACCATGTG LBb1.3 ATTTTGCCGATTTCGGAAC goxl2-1 LP TCGGTTCTGTTCGACCTTGG goxl2-1  goxl2-1 RP GTATAAACACGGATCGGATCC LBb1.3 ATTTTGCCGATTTCGGAAC goxl3-1 LP GATGATTTAGGAATAATATCTAGAGG goxl3-1  goxl3-1 RP TCCAGGATCATTGGTTTCGG LBb1.3 ATTTTGCCGATTTCGGAAC goxl4-1 LP2 TTCAAGCCAGTTTTTATAAAGATCG goxl4-1  goxl4-1 RP2 TTGGTGGAAGTAATCCTCACG LBb1.3 ATTTTGCCGATTTCGGAAC goxl4-2 LP2 TCATTTAATGGTGTCAGGATGC goxl4-2  goxl4-2 RP2 AGATCCACGCATAATGACTGG LB3 TAGCATCTGAATTTCATAACCAATCTCGATACAC goxl4-3 LP GAGCTTGGTATACCTGCATGC   goxl4-3  goxl4-3 RP ATATTACCCGATGGACGGATC   Spm32 TACGAATAAGAGCGTCCATTTTAGAGTGA goxl5-1 LP GCACCGTTAATGATCAAAACG goxl5-1  goxl5-1 RP CACCGTTAGCTATTCTCACCG LBb1.3 ATTTTGCCGATTTCGGAAC   40  Table 2.2: A list of primers used to genotype mutations used in this study. goxl5-2 LP GGGACTCTCTGTTCCCCC goxl5-2  goxl5-2 RP AGATTGAAAACGGATTCTCCC LB3 TAGCATCTGAATTTCATAACCAATCTCGATACAC goxl6-1 LP2 CAATGAAACAAACAGTGACTTGC goxl6-1  goxl6-1 RP2 CGACCTTAAAGAAAAGGGAGC LBb1.3 ATTTTGCCGATTTCGGAAC goxl6-2 LP GGATATGGTAAAGACCCTGCC goxl6-2  goxl6-2 RP GCTTTGTGTTCTGTGAAATTCG LB3 TAGCATCTGAATTTCATAACCAATCTCGATACAC goxl6-3 LP GGGGAGACGTCTTCTTACCAC goxl6-3  goxl6-3 RP GAGTTGCTTGCGTTGAGTACC Ds3-1 ACCCGACCGGATCGTATCGGT goxl6-3 LP GGGGAGACGTCTTCTTACCAC goxl6-4  goxl6-3 RP GAGTTGCTTGCGTTGAGTACC LB3 TAGCATCTGAATTTCATAACCAATCTCGATACAC  2.5 Mutagenesis, genetic analysis, and positional cloning of ruby Seeds homozygous for the mum2-1 mutation were mutagenised with ethylmethane sulphonate (Cat# M0880, Sigma). Seeds (100 mg; approximately 5000) were placed into 50 ml conical tubes and imbibed in 30 mL sterile water. EMS stock solution (120 μL) was added and seeds were left rotating gently overnight (16 to 17 hours). Seeds were then allowed to settle, and EMS solution was removed to a flask containing 100 mM sodium thiosulfate (Na2S2O3). Seeds were washed three times with 30 mL of 100 mM Na2S2O3, rotating gently for 15 min in Na2S2O3 each time. Seeds were similarly washed with 30 mL dH2O (15 min each), and then mixed into 400 mL 41  0.1% (w/v) agarose (cooled to RT). 20 – 50 seeds were plated onto solid AT medium to test germination rate. The remaining seeds were distributed 10 mL per pot into 40, 12 cm, round pots, keeping seeds mixed in agarose so that they were distributed evenly. M1 seeds for a single pot were harvested in bulk (each pot is one pooled M1 stock) and 70 – 100 M2 seeds from each M1 pool were planted for screening for suppression of the mum2 phenotype. M3 seeds were harvested individually from each M2 plant making sure to track from which M1 pool they originated. Seeds exhibiting suppression of the mum2 phenotype were individually selected from pools of mutagenised seeds and placed on AT medium. Seeds that germinated were transferred to soil and their seeds (M3) were tested to see if the suppression was a heritable trait, indicating a suppressor mutation in that line. To determine whether the mutations were recessive and in single nuclear genes, mum2-1 suppressor plants were crossed to mum2-1. The F2 generation of the cross was harvested and F3 seeds screened to see if the phenotypic segregation ratio resembles that of a single nuclear mutation (3 mum2:1 suppressor). Once confirmed to be recessive, suppressor lines were crossed to each other to determine the number of complementation groups. F2 seeds (harvested from F1 plants) from these crosses were stained with Ruthenium Red to determine if they exhibited a suppressor phenotype. If the phenotype was that of a suppressor rather than mum2, they were determined to be allelic. Crude-mapping of ruby was attempted by first crossing mum2-1 ruby-1 to Landsberg erecta (Ler) ecotype to generate plants that were heterozygous for Col-2 and Ler molecular markers. A segregating F2 population was genotyped for the mum2-1 mutation, and seeds of mum2-1 homozygous plants (F3 seeds) were screened for suppression. Only DNA of suppressors was used for positional cloning. Markers used for mapping are indicated in Table 2.3. Linkage 42  was inferred from molecular markers associated with the Col-2 ecotype that segregated with the suppressor phenotype. 43  Table 2.3: A list of primers used in crude-mapping. Chr Marker Marker position (Mb) BAC Forward primer (5ʹ-3ʹ) Reverse primer (5ʹ-3ʹ) I T7A14 1.4 T7A14 CTTTCTCTCAGTACGCAACCAG GGAATATTCACAAATGGCTCCTG I UPSC_I-2.579 2.5799 T23G18 CTCTTGGTGGTGTCCCAAGT TCGACGCAGTTTTTCATCAG I F16J7-TRB 3.8286 F16J7-TRB TGATGTTGAGATCTGTGTGCAG GTGTCTTGATACGCGTCGAT I F3F19-2 4.4 F3F19 CATATCTGCGTTAACGAATTTAGTAAT GGGCTAGGTTAGTCTCCCTTG I F14L17 4.8944 F14L17 AATACTATCTCAGCAGAAATGCAG AAACGAAAAGATAATGAAACTTTACCA I F10B6I-5 5.1514 F10B6I-5 TGTGCATGGTATTATAGGTGG AATCGCCTACTATATCTTTCAG I F9L1 5.2975 F9L1 AGAAATGAACAGGAGAATTGACTT TTTGACTCACTTTCACCACTTTG I F3O9 5.5 F3O9 GCCCTTCGTTTTTGTCGAT TTGAGGAACTTACAATTCTTGTCG I F20D23 5.8 F20D23 GCAATTTGAAGCGTTTTGTT GGTTTCCTTTTCAGGCAATTC I UPSC_I-6.975 6.9750 T20H2 CTTGAAACGTTATGTTATACTGCGC GAGAAAGAACAAGTGTGGACAAG I LUGSSLP887 7.4680 F10O3 ATTTTGGATTAACTTATGTTTATGCGT CATATACTGTCATAGTAAATGGTCCTTATCT I UPSC_I-8.660 8.6609 F21J9 GCGGCACAACCTAAATGAAA TGCATGCAATTATCACGTATG I CIW1 18.3639 F14J22 ACATTTTCTCAATCCTTACTC GAGAGCTTCTTTATTTGTGAT I UPSC_I-24.548 24.5490 F12P19 ACAAAATGCCGATCCAACAT TGCTGAAAACGTCAAGACCA II UPSC_II-1.401 1.4011 F3L12 GTTTGGATCAGTCCCAGCTC TGAAAAAGTGGTGGAACCAA II UPSC_II-5.794 5.7941 F17L24 TCATGCGGAAGTGAGTGTTC TGCTTGAGTTTGGTTTTTGC II CIW3 6.4031 T26I20 GAAACTCAATGAAATCCACTT TGAACTTGTTGTGAGCTTTGA II UPSC_II-12.560 12.5604 F16P2 CTCCAACACCACCTGCAA GAGATGGAGACCTGTTACGC II nga168 16.2920 T7F6 GAGGACATGTATAGGAGCCTCG TCGTCTACTGCACTGCCG  44  Table 2.3: A list of primers used in crude-mapping. III UPSC_III-3.058 3.0588 T22K18 GGATGCGAAATAAGCGATGA GGTGTAGCCGGCGTAAGTAA III UPSC_III-9.636 9.6366 MTC11 TTCAGCAACCTTCGATAAATCA CCATTGCCACCGTAGAAACT III UPSC_III-16.286 16.2862 T32N15 GGTTTGGTGGGAGAGAATGA CAAAAGAAATGCAACGAGACA III UPSC_III-2.2057 22.0577 T16L24 AATGCTTTGCATGCTTCAAT GAAGCAAGCACATGCCTAAA IV UPSC_IV-6.222 6.2223 F17A8 CAGAACCAAGCTGCAATGAA CCTTCGATGTCTTCGCTGAT IV UPSC_IV-11.840 11.8401 F7K2 ATTTACGGCGGTTCTTGATG TGCACCACACACATTCTCCT IV UPSC_IV-17.110 17.1105 F23E13 ATCGCTAACCCTCTCACGAA TGGCTGTGAGTGAGTGAAGA V CIW13 1.0067 F17C15/MED24 CGAACTTGAGACCTCTTGA GCTTACCTGGAGACAGTCA V MWD9V 7.3697 MWD9 GCAAGAAAAATGCTTACATGTT CAACGAAATCCAAATCCTCTC V CIW9 17.0440 K16E1/MFO20 CAGACGTATCAAATGACAAATG GACTACTGCTCAAACTATTCGG V UPSC_V-22.2317 22.3171 MBG8 GCATTGAAATAGTGTTTTTAACCAAA TGTTGGTTGCCACCTTATCA 45  After crude-mapping, a Next Generation Mapping approach was used to identify the mutation in the suppressor line, as described in Austin et al. (2011). Briefly, DNA was extracted from F4 seedlings, the progeny of 45 F3 lines exhibiting the suppressor phenotype by homogenising approximately 100 mg of tissue with a Precellys 24 homogeniser (Bertin Technologies Inc.) and 2 mm zirconium oxide beads (Next Advance Inc.) and extracting DNA using a PowerPlant Pro DNA Isolation Kit (Cat# 13400-50, MO BIO Laboratories). DNA was re-suspended in 10 mM Tris-HCl, pH 8 quantified using a NanoDrop 8000 spectrophotometer (Thermo Fisher Scientific), and equal amount of DNA (275 ng) from each sample was pooled. The solution was evaporated in a Savant DNA120 SpeedVac concentrator (Thermo Fisher Scientific) and re-suspended to 100 ng μL-1 in 10 mM Tris-HCl pH 8. Sequencing and analysis were performed at the Centre for the Analysis of Genome Evolution and Function at the University of Toronto, Toronto, ON, Canada (http://www.cagef.utoronto.ca/services/next-generation-genomics/). Candidate genes were identified using NGM online tool (http://bar.utoronto.ca/ngm/), and the identity of the RUBY gene was confirmed by Sanger sequencing all 5 candidate genes from lines homozygous for one of the other two alleles of ruby. In all three mutant lines, mutations were present only in AT1G19900. To confirm that RUBY is AT1G19900, additional T-DNA insertional mutants were ordered, and transgene complementation was done by transforming ProRUBY:RUBY-Citrine into mum2-1 ruby-1.  2.6 Molecular cloning and transgenic plants 2.6.1 ProRUBY:RUBY-Citrine RUBY (At1g19900) genomic DNA was amplified using Phusion High-Fidelity DNA polymerase (Thermo Fisher Scientific), including 4477 bp upstream of START codon, from Col-2 genomic 46  DNA using forward primer including an EcoRI restriction site (5′- GCTTAGAATTCGCTAACCCAATCTCAATCGAACC-3′), and reverse primer with an XbaI site (5′- ATAGTCTAGACCTCCTTCTAACTTTACCC -3′). The amplicon was gel-purified using EZ-10 Spin Column DNA Gel Extraction Kit (Bio Basic) according to manufacturer’s instructions, digested with EcoRI and XbaI, and purified using EZ-10 Spin Column PCR Products Purification Kit (Bio Basic). The resultant insert was ligated into de-phosphorylated pCambia2300 (Cambia) plasmid digested with EcoRI and XbaI, using T4 Ligase (NEB). Plasmid was cultured in DH5α E. coli strain (New England Biolabs) in liquid LB broth (Cat# 1.10285.5007, EMD Millipore) with 50 μg mL-1 kanamycin (Kan; Cat# K-120-25, Gold Biotechnology), purified using EZ-10 Spin Column Plasmid DNA Miniprep Kit (Bio Basic) and sequenced. For fluorescent tagging of RUBY, the Citrine-encoding sequence, together with a linker on its 5′ end and nosT on 3′ end, was amplified from pAD vector (DeBono, 2011) using forward primer with a SalI restriction site (5′- CTAGAGTCGACCCCTGGAGGTGGAGGTGGAGC-3′), and reverse primer containing an SbfI restriction site (5′- GCATGCCTGCAGGAGTAACATAGATGACACCGCGC-3′). The insert was subsequently sub-cloned into pCambia2300/ProRUBY-RUBY as described above. The final sequence-confirmed, binary vector was transformed into Agrobacterium tumefaciens strain GV3101 (pMP90). Resistant bacteria were selected on LB plates containing 50 μg mL-1 kanamycin (Kan), 25 μg mL-1 rifampicin (Rif; Cat# R-120-5, Gold Biotechnology), and 25 μg mL-1 gentamycin (Gent; Cat# G-400-10, Gold Biotechnology), at 28°C. A colony was grown overnight in 5 mL LB broth containing the same antibiotics, and sub-cultured in 200 mL LB broth with Kan/Gent overnight at 28°C. mum2-1 ruby-1 plants were transformed using the floral 47  dip method (Clough and Bent, 1998). T1 plants were selected on AT plates containing 35 μg mL-1 Kan, and positive transformants were transferred to soil.  2.6.2 ProRUBY:GOXL-Citrine Nopaline synthase terminator (nosT) was amplified from pGreenII 0229 using a forward primer with NotI restriction site (5ʹ-ctagagcggccgcGATCGTTCAAACATTTGGCAATAAAGT-3ʹ) and a reverse primer with SacI restriction site (5ʹ-agctggagctcGATCTAGTAACATAGATGACACCGCG-3ʹ), digested with NotI and SacI, and ligated using T4 ligase into de-phosphorylated pGreenII 0229 vector digested with the same enzymes. The vector was propagated in E. coli DH5α strain, isolated and sequenced. In the following step, ProRUBY was amplified from pCambia2300/ProRUBY-RUBY-Citrine vector using primers with KpnI restriction site (5ʹ- aattgggtaccGCTAACCCAATCTCAATCGAACCGG-3ʹ) and ApaI restriction site (5ʹ- agggggggcccTTTTTTGCTTTCAGGTTTAGTGTTTATTTG-3ʹ), and subcloned into pGreenII 0229/nosT. Citrine was then subcloned into the sequenced pGreenII 0229/ProRUBY:nosT after amplification with XbaI-Citrine-F (5ʹ-ctagttctagagCCTGGAGGTGGAGGTGG-3ʹ) and NotI-Citrine-R (5ʹ-cgatcgcggccgcTTATGCTAGAGGCGCAGCAGC-3ʹ) primers, and digestion with XbaI and NotI enzymes. After the final vector was propagated and sequenced, GOXL sequences from the transcriptional start site to the final base pair before STOP codon were amplified from genomic DNA of the Col-2 ecotype using primers listed in the Table 2.4. The amplicons and pGreenII 0229/ProRUBY:Cintrine:nosT  were digested with appropriate enzymes, ligated and propagated in E. coli and A. tumefaciens as described above. Plant transformation was performed by floral dipping of mum2-1 ruby-1 plants. 48  Table 2.4: A list of primers used to clone GOXL genes in-frame with Citrine. Primer 5ʹ-3ʹ Sequence ApaI-GOXL1-F aaaaagggcccATGAAAAAGTCAACAAGACTCTT HindIII-GOXL1- R atatcaagcttCCGACAATCTGAATCCATTCTCC ApaI-GOXL2-F aaaaagggcccATGGCAGAACTTATTCCTTG HindIII-GOXL2- R atatcaagcttCCAGAAACTATTCGTATCCATCG ApaI-GOXL3-F aaaaagggcccATGGCTGCAAAAGCCA SalI-GOXL3- R ataccgtcgacCCCTGTAAACGTACCCAAAGAC ApaI-GOXL4-F aaaaagggcccATGATCAACTCCAAAAATACATTCATCG HindIII-GOXL4-R atatcaagcttCCCTCAATCTGCACCCAAGC ApaI-GOXL5-F aaaaagggcccATGACGCAAGAAAGATTCAAGA HindIII-GOXL5-R atatcaagcttCCAATCTTAACCCAAACAGCTAC ApaI-GOXL6-F aaaaagggcccATGAAAGCCTCAACAAGAGTC HindIII-GOXL6-CC-R atatcaagcttCCGACGACCTGAATCCATTCTC  2.6.3 ProGOXL:GUS Non-coding regions upstream of transcriptional start sites of GOXL genes were amplified using primers listed in Table 2.4. Primers introduced restriction sites (Table 2.5, non-capitalised letters in sequences) that were digested using SalI and BamHI, or HindIII and BamHI restriction enzymes, and ligated into de-phosphorylated pBI101.1 plasmid digested with the same enzymes as amplicons (inserts). Plasmid propagation in E. coli and A. tumefaciens were performed as described above, and floral dip was done on Col-2 plants.  49  Table 2.5: A list of primers used to amplify putative promoter regions of GOXL genes. Primer name 5ʹ-3ʹ Sequence Amplicon length SalI-GOXL1p-F tgcaggtcgacTTATTGTCTCTATTAGAAAACTCTTGGG 488 bp BamHI-proGOXL1-R cccggggatccGATGGTTTGTCTAGTGAACATTGC SalI-GOXL2p-F tgcaggtcgacATGTAAAATATGATGACGCAAAC 1871 bp BamHI-proGOXL2-R cccggggatccACAGCCGTCGATTTATCTGTG SalI-GOXL3p-F tgcaggtcgacGAGTAAAACAATAATAACGGCGG 3987 bp BamHI-GOXL3p-R cccggggatccAAAAAAAAAAATTGTTGGTTGGAATTTTG HindIII-proGOXL4-F acgccaagcttGAGGAAGACCTTTTTTCAAAC 911 bp BamHI-proGOXL4-R cccggggatccAGTGGTTGGTGGTTTCTTCC HindIII-proGOXL5-F acgccaagcttAAAGATTTGAATCACCTGATTGC 626 bp BamHI-proGOXL5-R cccggggatccAAGTTGGTGAATTTTAGATCTG HindIII-proGOXL6-F acgccaagcttAGAAAGCCATGAAATTAGTGTAC 1125 bp BamHI-proGOXL6-R cccggggatccAGAACAAGCTCTTGTCTCTGC  2.6.4 CRISPR-Cas9 mutagenesis DNA sequences encoding guide RNAs (gRNAs) targeting GOXL1 and GOXL6 genes were amplified from pCBC-DT1T2 plasmid (Wang et al., 2015) using nested PCR with GOXL1-DT1-BsF (5ʹ-ATATATGGTCTCGATTGGATGACGATAATACTTCATGTT-3ʹ), GOXL1-DT1-F0 (5ʹ-TGGATGACGATAATACTTCATGTTTTAGAGCTAGAAATAGC-3ʹ), GOXL6-DT2-R0 (5ʹ-AACCTTACCACCCCAGGTTGTCCAATCTCTTAGTCGACTCTAC-3ʹ), and GOXL6-DT2-BsR (5ʹ-ATTATTGGTCTCGAAACCTTACCACCCCAGGTTGTCC-3ʹ), with gRNA sequences underlined. DT1-BsF/DT1-F0 and DT2-BsR/DT2-R0 primers were used in a 20:1 ratio. The insert was purified and digested using BsaI restriction enzyme and ligated into a BsaI-digested pHEE401E plasmid (Wang et al., 2015). Plasmid propagation in E. coli and A. tumefaciens were performed as described above, and floral dip was done on Col-2 plants. T1 50  plants were selected on AT medium with 30 μg mL-1 Hygromycin B. Putative mutant plants were selected for GOXL1 and GOXL6 sequencing based on apparent shorter siliques than Col-2.  2.7 Gene expression analyses 2.7.1 Reverse-transcription PCR (RT-PCR) A modified protocol based on Meisel et al. (2005) was used for total RNA extraction. Tissue (20-100 mg) was snap-frozen on dry ice, left at -70°C overnight, and ground to fine powder on dry ice using pre-chilled mortar and pestle, or with 2 mm zirconium oxide beads (Next Advance) on Precellys 24 tissue homogenizer (Bertin Technologies) at 6000 rpm for 20 s. Tissue was kept frozen until re-suspended in 500 μL of CTAB buffer pre-heated to 65°C, containing 2% (w/v) cetyltrimethylammonium bromide (CTAB, Cat# H5882, Sigma), 1.4 M NaCl (Cat# S271-3, Fisher Scientific), 20 mM ethylenediaminetetraacetic acid (EDTA, Cat# S311-100, Fisher Scientific), 100 mM Tris-HCl pH 8.0, 1% (w/v) polyvinylpyrolidone MW 40 000 (PVP40; Cat# 529504, Millipore Sigma), 0.05% (w/v) spermidine trihydrochloride (Cat# 85578, Sigma), and 5 mM DL-dithiothreitol (DTT; Cat# D0632, Sigma). Spermidine and DTT were added to the buffer right before use. Samples were mixed well on a vortex mixer and incubated at 65°C for 15 min. An equal volume (500 μL) of 24:1 (v/v) chloroform/isoamyl alcohol was added to samples, mixed on a vortex mixer, and centrifuged 12 000 g for 10 min at room temperature. The upper, aqueous phase was re-extracted one more time, and collected while carefully avoiding the interphase. RNA was precipitated overnight at -20°C by adding 10 M LiCl (Cat# L9650, Sigma) to 2 M final concentration. RNA was pelleted by centrifugation for 20 min at 16 000 g at 4°C, washed carefully in 75% (v/v) EtOH in DEPC-treated Milli-Q H2O, and centrifuged for 2 min at 16 000 g at 4°C. The wash was repeated once, EtOH removed, and pellet air-dried at room 51  temperature. Dry pellet was re-suspended in 10-15 μL DEPC-treated water, and total RNA quantified using a NanoDrop 8000 spectrophotometer (Thermo Fisher Scientific). The integrity of native RNA was inspected by agarose gel electrophoresis.  cDNA for RT-PCR was synthesised from 250 ng total RNA, treated with DNase I (Cat# 18068015, Thermo Fisher Scientific) according to manufacturer’s instructions. After inactivation of DNase, cDNA was synthesised using 5x All-in-One RT MasterMix (Cat# G490, Applied Biological Materials) according to manufacturer’s instructions in 10 μL final volume. For RT-PCR, 1 μL of cDNA was used in 10 μL reaction. ACT2 (At3g18780) and GAPC1 (At3g04120) were employed as reference genes. RT-PCR primers are described in the Table 2.6.  Table 2.6: A list of primers used in RT-PCR reactions. Primer name 5ʹ-3ʹ Sequence Gene amplified RUBY-RT-1R ACTTTACCCAAACACCTTCGC RUBY RUBY-RT-1F CGAATCTTCGTCCCAAGATTATATCTCC ACT2-F GTATTGCTCCTGAAGAGCACC ACT2 ACT2-R GACGGAGGATGGCATGAGG GAPC1-F TCAGACTCGAGAAAGCTGCTA GAPC1 GAPC1-R GATCAAGTCGACCACACGG  2.7.2 Histochemical promoter-GUS assays GUS activity was assayed using GUS buffer (1 mM 5-Bromo-4-Chloro-3-Indolyl-β-D-Glucoronide [X-Gluc], 100 mM NaPi pH 7, 10 mM EDTA, 0.5 mM potassium ferricyanide (K3[Fe(CN)6]), 0.5 mM potassium ferrocyanide (K4[Fe(CN)6]), 0.1% (v/v) Triton X-100). The buffer was made without X-Gluc and stored at -20°C (Stomp, 1992). GUS buffer was prepared 52  fresh each time by mixing 1 volume of fresh 20 mM X-Gluc in N,N-dimethylformamide and 19 volumes of buffer without X-Gluc. Organs assayed for GUS activity were pre-incubated in cold 90% (v/v) aqueous acetone on ice for 15-20 min and washed with GUS buffer without X-Gluc substrate. Samples were placed in fresh GUS buffer, vacuum infiltrated, and incubated at 37°C up to 24 h. Tissue was de-stained in EtOH/AcOH 1:1 (v/v), 70% (v/v) EtOH, and 95% (v/v) EtOH series. Samples were kept in EtOH at 4°C until imaged. Before imaging, samples were re-hydrated in 70% (v/v) EtOH, 50% (v/v) EtOH. and 30% (v/v) EtOH series. If necessary, samples were cleared in chloral hydrate/H2O/glycerol 8:3:1 (w/v/v) overnight (Willemsen et al., 1998).  2.8 Chemical analyses 2.8.1 Monosaccharide composition analysis using HPAEC-PAD For analysis of soluble mucilage, 20 mg of dry seeds hydrated in 1.4 mL of Milli-Q water or 20 mM Na2CO3, 5 μL of 5 mg mL-1 meso-erythritol (Cat# E7500, Sigma) was added as internal standard, and samples were shaken for 2 h. External standards for calibration were prepared by serial dilution (2 mM, 1 mM, 0.5 mM, 0.25 mM, 0.125 mM, 62.5 μM) from 100 mM monosaccharide mixture of L-fucose (Cat# F2252, Sigma), L-rhamnose (Cat# R3875, Sigma), L-arabinose (Cat# A3256), D-galactose (Cat# G6404, Sigma), D-glucose (Cat# G8270, Sigma), D-xylose (Cat# X3877, Sigma), D-mannose (Cat# M2069, Sigma), and D-galacturonic acid (Cat# Aldrich 85,728-9, Sigma). Standards were spiked with the same amount of meso-erythritol and processed the same way as the samples. 1 mL of mucilage extract was collected and dried under N2(g) at 60°C. Samples were incubated in 72% (w/v) H2SO4 on ice for 2 h to swell cellulose. Na2CO3 in samples was neutralised by adding additional H2SO4. After 2 h, H2SO4 was diluted to 4% w/v and sample volume brought to 0.5 mL with Milli-Q water. Samples were hydrolysed at 53  121°C at 15 psi for 1 h in an autoclave. Hydrolysates were filtered through 0.45 μm HV filters (Millipore) and loaded into HPLC vials. 15 μL samples were then injected using Spectra AS 3500 autoinjector (Spectra-Physics) onto a Dionex DX-600 high-performance liquid chromatograph (HPLC; Thermo Fisher Scientific) and separated using Dionex CarboPac PA1 column (Thermo Fisher Scientific). Neutral monosaccharides were separated at 1 mL min-1, with isocratic elution with water for 35 min, and sugars detected using a pulsed-amperometric detector with gold electrode after post-column addition of 200 mM NaOH at a flowrate of 0.5 mL min-1. The column was washed after each separation for 10 min with 250 mM NaOH and re-equilibrated with water. For acid sugars (GalA), the same samples were separated at 0.4 mL min-1, using linear gradient elution from 10 mM to 400 mM sodium acetate (NaOAc) for 30 min, and NaOH kept at constant concentration of 100 mM. The column was then washed for 10 min with 10 mM NaOAc, 300 mM NaOH and re-equilibrated with 10 mM NaOAc, 100 mM NaOH. Peak areas were integrated in Chromeleon software (Thermo Fisher Scientific), and data processed using Python 2.7 with NumPy and SciPy packages.  For whole seed analysis, 20 mg of seed was frozen on dry ice, then ground on dry ice using mortar and pestle. Alternatively, frozen seeds were ground with 2 mm zirconium oxide beads (Next Advance Inc.) on Precellys 24 tissue homogeniser (Bertin Technologies Inc.) at 6000 rpm for 20 s. Samples were suspended in 1 mL 70% (v/v) aqueous EtOH, incubated for 10 min at 65°C, vortexed for 10 min at RT, and centrifuged at 16000 g for 30 s. Washed in 70% (v/v) EtOH, followed by 10 min vortex and centrifugation was repeated once. Three more washes were completed to prepare the final alcohol-insoluble residue (AIR): 80% (v/v) MeOH, 100% MeOH and acetone. Acetone was finally evaporated under a gentle stream of N2(g), and the AIR weighed and transferred to glass tubes for hydrolysis. 20 μL of 5 mg mL-1 meso-erythritol 54  was added to samples, dried under N2(g), and processed and analysed similar to that of the mucilage samples (above). The final volume of H2SO4 was adjusted so that, once diluted, the final concentration was 4% (w/v) H2SO4 in 2 mL. For sequential extraction of mucilage, outer non-adherent mucilage was extracted from 100 mg of intact seeds with water (3 mL) for 3 h at room temperature. After centrifugation (8 000 g, 5 min), supernatants were carefully removed, filtered through a disposable glass microfiber filter (13 mm diameter, 2.7 µm pore size; Whatman) and retained for analysis. Seeds were then rinsed with 5 mL of 50 mM sodium acetate buffer, pH 4.5 (three changes) and rhamnogalacturonan hydrolase (UniProt Q00018; Novozymes, Copenhagen, Denmark) was added to the washed seeds (0.8 nkat). The inner adherent mucilage extracts were recovered as described previously (Sullivan et al., 2011). Seeds following mucilage removal were carefully rinsed with distilled water (three changes), freeze-dried, and ground with mortar and pestle. They were further delipidated with 2 mL methanol/chloroform (1/2; v/v) overnight with head-over-tail mixing at room temperature, centrifuged (8 000g, 10 min), and supernatant removed. The treatment was repeated twice with 2 h of head-over-tail mixing, after which the samples were dried overnight at 40°C. Uronic acid (as GalA) was determined by automated m-hydroxybiphenyl method (Thibault, 1979) either directly (mucilage extracts) or after pre-hydrolysis (H2SO4 72%, 30 min at RT) and hydrolysis (H2SO4 2 N, 6h at 100°C) for demucilaged, delipidated seeds. Individual neutral sugars were analysed as their alditol acetate derivatives (Blakeney et al., 1983) by gas-liquid chromatography after hydrolysis with 2 M trifluoroacetic acid at 121°C for 2.5 h for mucilage extracts or pre-hydrolysis (H2SO4 72%, 30 min at room temperature) and hydrolysis (H2SO4 2 N, 6h at 100°C) for demucilaged delipidated seeds. 55   2.8.2 Per-O-methylation and Linkage analysis of neutral sugars Mucilage was prepared for PMAA glycosyl linkage analysis by shaking 20 mg of Col-2 and ruby-1 seeds in 25 mM Na2CO3 for 2 h. The seeds were left to settle, and the supernatant was removed and dried under N2(g) at 60 °C. Dry mucilage was sent to the Complex Carbohydrate Research Center, University of Georgia, Athens, GA, USA for analysis. The analysis was done on biological duplicates, as described by York et al. (1986). Initially, dry sample was suspended in about 300 μL of dimethyl sulphoxide (DMSO) and placed on a magnetic stirrer for 1 week. The sample was  permethylated according to Ciucanu and Kerek (1984). Solid NaOH was added to the sample and incubated for 15 min at room temperature, followed by the addition of methyl iodide (CH3I) and 45 min incubation. Additional NaOH was added, followed by 10 min incubation, and finally more CH3I was added and samples incubated for 40 min. Following the derivatisation, the permethylated material was hydrolysed using 2 M trifluoroacetic acid (TFA) for 2 h in a sealed tube at 121°C, reduced with NaBD4, and acetylated using acetic anhydride/TFA. The resulting permethylated alditol acetates (PMAAs) were analysed on a Hewlett Packard/Agilent 7890A GC gas chromatograph (Agilent Technologies Inc.) interfaced to a 5975C MSD (mass selective detector, electron impact ionization mode EI-MS; Agilent Technologies Inc.); separation was performed on a 30 m Supelco SP-2380 bonded phase fused silica capillary column (Sigma-Aldrich Co.).  2.8.3 Digestion and analysis of mucilage RG-I For the analysis of Gal oxidation in the mucilage, outer soluble mucilage was extracted from WT (Col-2), ruby, mum2 and ruby mum2 seeds (100 mg) with 0.05 M HCl (5 mL) for 30 min at 56  85°C, and then with 5 mL of 0.3 M NaOH as previously described (Macquet et al., 2007b). Seeds were rinsed 3 times with 5 mL of water, acidified to pH 4.5 with 0.05 M HCl, and then rinsed with 5 mL of 50 mM sodium acetate buffer, pH 4.5 (three changes). Rhamnogalacturonan hydrolase (UniProt Q00018; Novozymes, Copenhagen, Denmark) was added to washed seeds (0.8 nkat) and inner adherent mucilage extracts were recovered as described previously (Sullivan et al., 2011). The oligosaccharides were analysed using HPAEC-PAD as described by Ralet et al. (2010) and using IP-RP-UHPLC-MS/MS method described below.  2.8.4 IP-RP-UHPLC-MS/MS Chromatographic separation was achieved on an Ultra High-Performance Liquid Chromatography system (UHPLC, Acquity H-Class® Waters, Manchester, UK), fit with a BEH C18 column (100 mm × 1 mm, packed with 1.7 µm porosity particles; Waters, Manchester, UK). A ternary gradient was used (A: Milli-Q water, B: 100% methanol and C: 20 mM heptylammonium formate, pH 6), from 2% to 25% of solvent B in 10 min, then up to 73% at 23.5 min and maintained at 73% for 4 min. Percentage of solvent C was kept constant at 25%. The flow rate was of 0.15 mL min−1 and column was heated to 45°C. Mass spectrometry measurements were performed using a Synapt G2 Si HDMS (Waters, Manchester, UK) in negative ionisation mode and in sensitivity TOF measurements mode. The parameters used for electrospray were the following: capillary voltage: 2.5kV; sampling cone: 50; desolvation temperature 250 °C; desolvation gas: 350 L/h. MS/MS measurements were performed in the trap cell of the triwave cell using an energy of 50. 57  2.8.5 Extraction and analysis of seed surface phenolics To extract ester-linked hydroxycinnamic acids from the mucilage and cell walls of the seed surface, 20 mg of wild type and mutant seeds (exact mass recorded) were hydrated in 1 mL of de-gassed 2 M NaOH. 20μL of 5 mg mL-1 vanillin (Cat# V2375, Sigma) was added as an internal standard, samples were flushed with N2(g) and incubated at 30°C for 2 h. Liquid was transferred to glass tubes, acidified with 150 μL 72% (w/v) H2SO4, and extracted 3x with 1 mL of ethyl acetate (EtOAc). All three EtOAc extracts were combined and evaporated under N2(g). Samples were re-suspended in 0.5 mL of 50% (v/v) aqueous acetonitrile (MeCN), 0.1% (v/v) TFA, filtered through 0.45μm HV filters, and loaded into plastic HPLC vials. Sinapic acid (Cat# D13,460-0, Aldrich Chem. Co.) standards were made by serial dilution from 50 mM stock in MeOH, spiked with the same amount of vanillin as the samples, evaporated under N2(g), and re-suspended in 50% (v/v) MeCN, 0.1% (v/v) TFA. 15 μL of sample was injected by an ASI-100 autoinjector (Dionex) and separated on Symmetry C18 column (5 μm, 4.6 mm x 250 mm; Waters) at 35°C on a Summit® HPLC system (Dionex) fit with a PDA-100 Photodiode Array Detector (Dionex). H2O with 0.1% (v/v) TFA (solvent A) and MeCN/MeOH 3:1 with 0.1% (v/v) TFA (solvent B) were used for separation at a flow rate of 0.7 mL min-1 using linear gradient from 5% to 45% solvent B, 0-40 min. Analytes were detected by measuring absorbance at 255 nm and 320 nm. The column was washed for 10 min with 75% solvent B, and re-equilibrated for 20 min with 5% solvent B.  For LC-ESI-MS identification of seed surface phenolics, Col-2 and ruby-1 samples were processed as described above and delivered dry under N2(g) to the Mass Spectrometry Services, Department of Chemistry, Simon Fraser University, Burnaby, BC, Canada (https://www.sfu.ca/chemistry/research/facilities/massspec.html). Samples were analysed on 58  Agilent 1200 SL LC system (Agilent Technologies). They were separated on a Zorbax XDB C18 column (1.8 μm, 50 mm x 4.6 mm; Agilent Technologies), at 30°C, with a flowrate of 0.6 mL min-1. H2O with 0.1% (v/v) formic acid (Solvent A) and MeCN with 0.1% (v/v) formic acid (Solvent B) were used for separation as follows: 10% B at 0 min, 10% B at 1min, 40% B at 8 min, 100% B at 9 min, 100% B at 10 min, 10% B at 10.1 min, 10% B at 12 min (stop). Analytes were detected using maXis Impact Ultra-High Resolution tandem TOF (UHR-Qq-TOF) mass spectrometer (Bruker) in 50 – 1500 Da mass range (ESI in negative mode; source capillary 4.5 kV; gas temperature 200°C; gas flow 9 L min-1).  2.9 Phylogenetic analysis Amino acid sequences used in phylogenetic analyses were obtained from UniProt (Bateman et al., 2017) for characterised members of AA5 CAZy family of enzymes, or from Plaza 4.0 for all the plant proteins (Van Bel et al., 2018). Based on initial multiple alignment of sequences, the ends of sequences were truncated in Mesquite 3.51 (http://www.mesquiteproject.org) to remove components outside the catalytic domain. Truncated sequences were aligned using MAFFT algorithm (Katoh and Standley, 2013), and alignment visualised using JalView (Waterhouse et al., 2009). Datasets were tested for appropriate protein evolution models using ProtTest 3.4.2 (Darriba et al., 2011). Alignments were used to construct phylogenetic trees using raxmlGUI (Silvestro and Michalak, 2012), running RAxML 8.1.2 (Stamatakis, 2014) using amino acid substitution models suggested by ProtTest, and 1000 replications to generate bootstrap branch support values. Unrooted tree was visualised in FigTree v1.4 (http://tree.bio.ed.ac.uk/software/figtree/). 59  2.10 Protein expression and purification 2.10.1 Protein expression and purification in E. coli RUBY was cloned with and without predicted signal sequence, based on a SignalP 4.1 prediction. The gene was amplified from pCambia2300/ProRUBY:RUBY-Citrine:nosT vector together with 3′ linker sequence using a forward primer with a SacI restriction site for both full length (5′-aattcgagctccggaggtatggcagcacgagcaacattc-3′) and sequence without the signal sequence (5′-aattcgagctccggaggtgcccgaggattatggaaatacatcg-3′). The same reverse primer was used to amplify both inserts, and it contained a XhoI site (5′-tggtgctcgagtccacctccacctccaggg-3′). After digestion and purification, the inserts were ligated into the pET-21b (+) vector (Novagen) in-frame with the C-terminal His6-tag. The plasmid was then cultured in DH5α, purified and sequenced. For soluble protein expression, plasmids were transformed into Arctic Express (DE3) E. coli (Agilent Technologies). Expression was assessed on a small scale, and full-length protein was found to be poorly expressed, thus protein without the signal peptide was used in purification. For purification, starter culture was grown overnight at 37°C in LB broth with 100 μg mL-1 Carbenicillin (Carb; Cat# BP2648, Fisher Scientific) and 25 μg mL-1 Gent. The next day, cells were sub-cultured in 500 mL TB medium (2.4% yeast extract, 1.2% tryptone, 0.4% glycerol, 100 mM KPi buffer pH 7.5) with 50 μg mL-1 Carb. Culture was grown in 2 L flask at 37°C at 250 rpm until an O.D. 600 of 0.8-1.2 was achieved. Expression was induced by adding Isopropyl-β-D-thiogalactopyranoside (IPTG; Cat# 10724815001, Roche) to 0.1 mM. Protein was expressed at 11°C for 36 h, after which the culture was centrifuged for 20 min at 10 000 g. The pellet was re-suspended in 35 mL of extraction buffer (100 mM NaPi buffer pH 7.4, 10% [w/v] sucrose, 500 mM NaCl, 0.3 mM PMSF), lysozyme (Cat# L6876, Sigma) was added at 0.5 mg mL-1, and the suspension incubated on ice for 1 h. Cells were lysed using Vibra Cell™ VC505 sonicator fit 60  with a CV334 converter and 13 mm tip probe (Sonics and Materials, Inc.) at 40% amplitude, for 15 min (5 s ON, 5 s OFF). The lysate was brought to 50 mL with cold buffer and centrifuged for 25 min at 12000 g and 4°C. The supernatant was loaded on equilibrated HisPur Ni-NTA agarose resin (Thermo Fisher Scientific) in a glass column, and resin with sample was incubated for 1 h at 4°C, after which the flow-through was collected, and resin washed 2x with 10 mL of 100 mM NaPi  buffer pH 7.4, 10% (w/v) sucrose, 500 mM NaCl, 30 mM imidazole, then 2x with 10 mL of 100 mM NaPi buffer pH 7.4, 10% (w/v) sucrose, 500 mM NaCl, 30 mM imidazole, 5 mM ATP, 5 mM MgCl2, followed by 3 washes with 10 mL 100 mM NaPi buffer pH 7.4, 10% (w/v) sucrose, 500 mM NaCl. Protein was eluted 5 times with 2 mL 100mM NaPi buffer pH 7.4, 10% (w/v) sucrose, 500 mM NaCl, 250 mM imidazole. A280 of eluted fractions was determined using a NanoDrop 8000 spectrophotometer (Thermo Fisher Scientific), and fractions with highest reads were combined and desalted in 100 mM NaPi pH 6.0, 10% (w/v) sucrose, 500 mM NaCl on Econo-Pac 10DG column (Bio Rad) at 4°C using minimal dilution protocol. For activation, 0.5 mM CuSO4 was added to the buffer, as described by Spadiut et al. (2010), and mixed on a head-over-tail mixer at 4°C overnight. Proteins were analysed using SDS-PAGE and Western blot with His-Probe (H-3) antibody (Santa Cruz Biotechnology). For activity assays, cells carrying the pET-21b empty vector were grown under the same conditions alongside cultures with pET-21b/RUBY-His6 and used as a negative control in activity assays. Using buffers described in the section below, and galactose and glycerol as substrates, activity was assayed on crude lysates, supernatants, and all the fractions derived from the purification. 61  2.11 Enzyme activity experiments 2.11.1 Enzyme assays with seeds For substrate screening using dry seeds, ~50 seeds (Col-2 and ruby-1) were incubated in 100 μL of 50 mM NaPi buffer pH 6.0, 2 U mL-1 Horseradish Peroxidase, Type II (HRP; Cat# P8250, Sigma), 100 μM 3,3',5,5'-tetramethylbenzidine (TMB; Cat# 229280010, Acros Organics), and 10 mM substrate. Samples were incubated overnight at RT protected from light. A compound was considered a substrate if a blue colour developed in Col-2, but not ruby-1 samples. The effects of activators and inactivators were assayed on 3 mg of seeds (exact mass recorded) of 3 biological replicates of Col-2 seeds. Seeds were pre-incubated for 1 h in 1 mL Milli-Q water at RT, washed twice, and incubated for 1 h in Milli-Q water at RT (control), 1 mg mL-1 Proteinase K, 1 mM CuSO4, 1 mM EDTA, or Milli-Q water at 95°C (heat inactivation). After 1 h pre-incubation, the samples were washed 2x in 2 mL 200 mM NaPi buffer pH 6.0, and Proteinase K inhibited at 65°C for 10 min. Activity was assayed in 2 mL of 50 mM NaPi buffer pH 6.0, 2 U mL-1 HRP, 100 μM TMB, 200 mM D-galactose at RT (22°C). Seeds were mixed by inverting, and 150 μL were sampled at multiple timepoints once the colour started to develop. Sampling was done within linear range (blue colour). Immediately before reading samples at A450 on a Synergy HT microplate reader (BioTek), 10 μL of 1 M H2SO4 were added to each sample to stop reactions. H2O2 in the solutions was calculated based on calibration curves of standards with known concentrations of H2O2 within a linear range of detection. Standard solutions were loaded on the same microtiter plates in the same volumes as the samples, and values calculated based on Beer-Lambert law. For calculation of specific activity (nmol min-1 mg-1 seeds), only timepoints that showed linearity were used. 62  Specific activity was assayed on 3 mg of seeds (exact mass recorded) of 3 biological replicates, using Col-2, ruby-1 and ruby-5 genotypes. Seeds were pre-incubated in Milli-Q water for 2 h at RT, washed twice, and left in 1 mL 2x reaction buffer (100 mM NaPi buffer pH 6.0, 4 U mL-1 HRP, 200 μM TMB). Reactions were initiated by addition of 1 mL of 2x substrates in Milli-Q water to give final substrate concentrations of 200 mM (D-galactose, raffinose, meso-erythritol and glycerol) or 40 mM (ONP-β-D-galactose). Data were collected and processed the same way as for activation and inactivation assays. Qualitative enzyme assays were performed on mum2-1 ruby-1 seeds expressing ProRUBY:GOXL by pre-hydrating seeds for 1.5 h in 100 mM KHCO3 freshly diluted from 500 mM stock solution. Seeds were washed twice in 50 mM buffer used in the assay: NaOAc buffer (pH 4.5, pH 5.0, pH 5.5) and NaPi buffer (pH 6.0). 2x reaction buffer (4 U mL-1 HRP, 200 μM TMB, 800 mM D-galactose) was added to 100 mM buffer to start the reaction. Once the blue colour was developed, the plate was scanned.  2.11.2 Detection of galactose oxidase substrates in the mucilage Col-2, mum2-1, ruby-1 and mum2-1 ruby-1 seeds (200 mg; 3 biological replicates) were shaken in 5 mL of 20 mM Na2CO3 for 2 hours at room temperature. After the seeds settled at the bottom, the supernatant was collected, and seeds washed in 5 mL of Milli-Q water. Both supernatants were pooled, then separated into two samples of equal volumes. NaBH4 was added to 0.1% (w/v) from a 1% (w/v) stock in 0.2 M NaOH, and the other sample was left untreated (control). After 15 min, the samples were neutralised by adding glacial acetic acid, precipitated by adding 4 volumes of 95% (v/v) EtOH, and pelleted by 10 min centrifugation at 7000 g. Supernatants were aspirated, pellets re-suspended in Milli-Q water, and dialysed in cellulose acetate tubing (12-14 63  kDa MWCO) against Milli-Q water. Samples were then frozen on dry ice and lyophilised using a FreeZone 4.5 L freeze dry system (Labconco). Dry mucilage was weighed and re-suspended to 2 mg mL-1 in Milli-Q water. Samples were mixed on a vortex mixer for 2 h at RT to enhance re-suspension and stored at 4°C until the analysis.  Extracted mucilage was imaged for opacity/transparency by loading 150 μL of 2 mg mL-1 mucilage samples into 96-well microtiter plate and taking images with Nikon Coolpix 8400 digital camera. The effect of Na2CO3 on mum2 mucilage solubility was examined by splitting 2 mg mL-1 mum2-1 mucilage samples into two 140 μL samples. They were loaded into a 96-well plate, 10 μL of H2O was added to one (control) and 10 μL of 0.5 M Na2CO3 to the other (Na2CO3). The plate was incubated at RT for 10 min and imaged the same as above. Three biological replicates were used.  Availability of galactose as a substrate was measured by adding 50 μg of mucilage in buffer containing 50 mM NaPi pH 6.0, 100 μM TMB, 1 U mL-1 HRP and 1 U mL-1 galactose oxidase (GalOx, Cat# G7400, Sigma) to a final volume of 150 μL in 96-well polystyrene microtiter plate. The plate was left in the dark at room temperature for 1 h, reactions stopped by adding 10 μL of 1 M H2SO4, and absorbance at 450 nm measured within 10 min using a Synergy HT microplate reader (BioTek). The concentration of H2O2 was measured using standard solutions with the same volumes as the samples.  2.11.3 Mucilage insolubility experiments For reduction at neutral pH, 50 – 100 seeds were mixed with 900 μL of 1 M imidazole-HCl, pH 7.0. After 30 min incubation in the buffer, 100 μL of 1% (w/v) NaBH4 in 10 mM NaOH (10 mM NaOH to control samples). Seeds were mixed on a head-over-tail mixer for 2 h at room 64  temperature, after which the pH was measured using pH indicator strips to ensure that it remained stable. Seeds were washed twice with H2O, once in 1% (w/v) aqueous glacial acetic acid, twice in H2O, stained with 0.02% (w/v) ruthenium red, and imaged.  Dehydration – re-hydration experiments were completed as follows: mucilage was released by mixing 50-100 seeds in 1425 μL of 20 mM Na2CO3 for 2 h at room temperature. 75 μL of 1% (w/v) NaBH4 in 0.2 M NaOH (0.2 M NaOH only to control samples) was added for reduction, and samples were mixed for another hour. Seeds were washed twice with H2O, once in 1% (w/v) aqueous glacial acetic acid, twice in H2O, vacuum-filtered, and dried with filter papers on silica beads overnight. The next day, seeds were collected into plastic tubes, hydrated in H2O, mixed for 2 h at room temperature, stained with 0.02% (w/v) ruthenium red, and imaged. 65  Chapter 3: RUBY PARTICLES IN MUCILAGE (RUBY) is a putative galactose oxidase that cross-links pectin 3.1 Introduction The emergence of multicellularity necessitated the development of mechanisms that promote cell-to-cell adhesion. In plants, cell adhesion is mediated largely through the middle lamella, an extracellular matrix rich in pectins and structural proteins that is shared between the walls of two adjacent cells (Zamil and Geitmann, 2017). Three major types of pectic polysaccharides have been described: homogalacturonan (HG), rhamnogalacturonan I (RG-I), and rhamnogalacturonan II (RGII). HG has the simplest structure consisting of α-(1→4)-linked galacturonic acid (GalA) monosaccharides (McNeil et al., 1984). In contrast, RG-I has a backbone composed of alternating rhamnose (Rha) and GalA monosaccharides linked as [→2)-α-L-Rhap-(1→4)-α-D-GalpA-(1→] (McNeil et al., 1980). In addition, the rhamnose can act as branch points for diverse oligosaccharide side-chains composed of arabinose (Ara) and/or galactose (Gal) (Lau et al., 1987; Lerouge et al., 1993). RG-II has a backbone that resembles HG, but its side-chains are highly diverse, consisting of 12 monosaccharides linked in a specific manner which is conserved among many plant species (O’Neill et al., 2004). Immunolabelling experiments indicate that middle lamellae between cells contain HG, RG-I and hydroxyproline-rich glycoproteins (HRGPs; Bush et al., 2001; Moore et al., 1986; Smallwood et al., 1994; Willats et al., 2001). Mutants affecting cell adhesion have defects in one of the three pectic polysaccharides. Defects in biosynthesis of HG in quasimodo1 (qua1) and quasimodo2 (qua2) mutants result in loss of cell adhesion in roots, hypocotyls, and leaves (Bouton et al., 2002; Mouille et al., 2007). Similarly, when an HG-degrading polygalacturonase is ectopically expressed in apples, the result 66  is a loss of cell adhesion (Atkinson et al., 2002). Mutants for the putative polygalacturonase gene of Arabidopsis, QUARTET3, and its functional partner, the pectin methylesterase QUARTET1, lack pollen tetrad separation which requires HG degradation (Francis et al., 2006; Rhee et al., 2003). A mutant affecting cell adhesion of Nicotiana plumbaginifolia callus, nolac-H18, lacks a portion of the RG-II side-chain, which prevents RG-II cross-linking through borate ions (Iwai et al., 2002). A number of studies have implicated the involvement of arabinans and galactans in cell-to-cell adhesion. For example, the N. plumbaginifolia cell adhesion mutant nolac-H14 lacks arabinan side-chains (Iwai et al., 2001). The tomato Cnr mutant has a cell adhesion phenotype and a change in arabinan distribution in the cell walls of fruit pericarp (Orfila et al., 2001). Arabidopsis plants deficient in FRIABLE1, a putative O-fucosyltransferase, exhibit cell adhesion and organ fusion phenotypes, as well as changes in arabinose and galactose-containing oligosaccharides in the Golgi apparatus (Neumetzler et al., 2012). A hallmark of fruit softening in many species is the removal of arabinan and galactan side-chains. For example, the softening of nectarines (Prunus persica) is correlated with the degradation of RG-I-associated galactans and solubilisation of pectins rich in arabinans (Dawson et al., 1992). In apples, these side-chains were specifically assigned to RG-I (Peña and Carpita, 2004). In ripe carambola fruit, β-galactosidase was found to be involved in the removal of galactans, and responsible for solubilisation of pectins and tissue softening (Balasubramaniam et al., 2005). Despite this evidence, the mechanism(s) through which arabinans and galactans associated with RG-I and cell wall structural proteins influence cell-to-cell adhesion remain unclear. Due to high RG-I abundance and available mutants, Arabidopsis seed coat mucilage was used as a tool to investigate the role of galactans and arabinans in pectin cohesion. In this chapter, a forward genetic approach was undertaken to find suppressors of the mum2 phenotype. 67  One suppressor, ruby particles in mucilage (ruby), affects mucilage properties of mum2, as well as cell adhesion in the seed coat epidermis. RUBY encodes a putative galactose oxidase that appears to strengthen the middle lamellae and mucilage by cross-linking branched RG-I. These data reveal a mechanism of cross-linking within the middle lamellae, provide strong evidence for a biological role for plant galactose oxidases, and demonstrate the importance of arabinogalactan side chains and oxidation in cell wall biology.  3.2 Results 3.2.1 Mutations in RUBY can suppress the mum2 mucilage extrusion phenotype To investigate the mechanism by which RG-I side chains influence mucilage extrusion, a genetic modifier screen to find suppressor mutations of mum2 was employed. A population of mum2-1 seeds was mutagenized with ethyl methanesulfonate (EMS) and M3 seeds from individual M2 plants were screened for wild type-like mucilage extrusion when exposed to water. From 2469 M2 lines screened, 3 lines extruded a mucilage capsule similar to wild type, but with small particles that stained dark red when treated with ruthenium red (Figures 3.1 C, D, G and H). Based on phenotypic ratios in a cross of the ruby-1 mum2-1 double mutant to mum2-1, ruby-1 segregated as a single nuclear recessive mutation (3 mum2:1 suppressor; Χ2= 0.0045662, df = 1, p = 0.9461, n = 73). Allelism tests confirmed that all three mutants were homozygous for mutant alleles of the same gene (Figures 3.1 I to K). Based on the novel phenotype (Figure 3.1 D, arrowheads), we named this gene RUBY PARTICLES IN MUCILAGE (RUBY), and accordingly the mutant alleles ruby-1, ruby-2 and ruby-3. To determine the mucilage phenotype of the ruby single mutants, ruby single mutants were isolated from the F2 of crosses between ruby mum2-1 double mutants and wild type. The extruded mucilage of ruby-1 and ruby-3 was similar to that of 68  mum2-1 ruby-1 and mum2-1 ruby-3, respectively (compare Figures 3.1 C and D, and G and H) in all aspects of the observed phenotypes whereas ruby-2 resembled wild type more than mum2-1 ruby-2 (compare Figures 3.1 E and F). In addition, mum2-1 ruby-1 and mum2-1 ruby-3 (Figures 3.1 C and G), demonstrated stronger suppression than mum2-1 ruby-2 (Figure 3.1 E). Further phenotypic characterisation was done on ruby-1.   69   Figure 3.1: ruby exhibits multiple seed coat mucilage phenotypes. (A) to (M) seeds agitated in water for 2 h and stained with Ruthenium Red. Bars = 200 μm. (A) to (H) Three suppressor lines homozygous for an allele of ruby and mum2. (A) Wild type (Col-2), (B) mum2-1, (C) mum2-1 ruby-1, (D) ruby-1, (E) mum2-1 ruby-2, (F) ruby-2, (G) mum2-1 ruby-3, (H) ruby-3. Black arrowheads indicate ruby particles in mucilage. Black arrows indicate primary cell wall being released in sheets. (I) to (K) F2 seeds harvested from individual F1 plants resulting from crosses between different suppressor lines. (I) mum2-1 ruby-1 x mum2-1 ruby-2 cross, (J) 70  mum2-1 ruby-1 x mum2-1 ruby-3 cross, (K) mum2-1 ruby-3 x mum2-1 ruby-2 cross. Note that all pairwise crosses failed to complement each other for the ruby suppression (L) and (M) ruby can suppress bxl1-1. (L) bxl1-1, (M) bxl1-1 ruby-1. (N) and (O) Non-stained (N) wild type and (O) ruby-1 seeds imaged using DIC microscopy after shaking in water. White arrow indicates columella without primary cell wall attached in the mutant. White arrowhead indicates primary cell wall attached to the top of columella in wild type and detached in the mutant. Bars = 20 μm. (P) Box-plot showing surfaces of adherent mucilage halo for Wild type (Col-2) and ruby-1. Asterisks indicate p < 0.001 based on Welch’s t-test. n = 130. (Q) and (R) seeds hydrated and agitated in water, followed by staining with Pontamine Fast Scarlet 4B. (Q) Wild type (Col-2), (R) ruby-1. Arrows indicate cellulosic rays. Bars = 50 μm.  3.2.2 RUBY is required for mucilage integrity Aside from the strong suppression of the mum2 extrusion phenotype and the presence of ‘ruby’ particles, the extruded mucilage of ruby seeds possessed several other phenotypes that could be identified by light microscopy. First, the ruby adherent mucilage halo was not smooth like wild type, but had rough edges giving the halo a dishevelled appearance (Figures 3.1 C and D; arrows). Second, the outer primary cell walls of ruby epidermal cells frequently detached from seeds (Figure 3.1 O, arrowhead), instead of remaining attached to the top of columellae (Figure 3.1 N). Third, the adherent mucilage halo appeared larger than that of wild type. To confirm this observation, the area occupied by the adherent mucilage of Ruthenium Red-stained seeds was measured as described by Voiniciuc et al., (2015). Based on pair-wise comparison using two-tailed Welch’s t-test, the ruby halo is significantly larger than that of wild type (Figure 3.1 P). These data indicate that RUBY may have a role in making the mucilage halo more compact. Fourth, the extrusion dynamics of ruby to wild type were compared by exposing mature dry seeds to 0.02% ruthenium red and filming mucilage extrusion. Wild-type seeds readily extruded 71  mucilage within seconds and the non-adherent halo expanded rapidly (Figure 3.2 A). Compared with the non-extruding mum2 (Figure 3.1 B; Figure 3.2 B), the ruby mutant appears to extrude at a similar rate to wild type (Figure 3.2 D), but the radial cell wall rupturing during the extrusion did not appear wild type-like. Fifth, when seeds are stained for cellulose with Pontamine Fast Scarlet 4B, wild-type seeds show prominent rays attached to the surface of columellae (Figure 3.1 Q, arrow; Griffiths et al., 2014). In contrast, ruby-1 mutant seeds have what seems to be a collapsed cellulosic ray above the columellae, changing the shape of rays from rod-like to conical (Figure 3.1 R, arrow). This implicates RUBY in the organisation of mucilage cellulose. To understand if the ability of the ruby mutation to suppress mucilage extrusion defects is specific to mum2, ruby was crossed to bxl1. The bxl1-1 mutant exhibits patchy, reduced, mucilage extrusion (Figure 3.1 L) and a delay in the extrusion (Figure 3.2 E), as well as an increase in terminal arabinose residues on RG-I (Arsovski et al., 2009). The bxl1-1 ruby-1 double mutant extruded mucilage at a higher degree than bxl1-1 (Figure 3.1 M; Figure 3.2 F), demonstrating that ruby is a suppressor of bxl1-1. These data show that the ability of ruby mutations to suppress the loss of mucilage expansion is not specific to mum2 mucilage.     72   Figure 3.2: ruby suppresses mum2 and bxl1 phenotypes to a different degree. Images representing 4 timepoints extracted from mucilage extrusion movies. Mucilage extrusion of a single seed was filmed in 0.02% Ruthenium Red. (A) Wild type (Col-2); (B) mum2-1; (C) mum2-1 ruby-1; (D) ruby-1; (E) bxl1-1; (F) bxl1-1 ruby-1. 73  3.2.3 RUBY is required for cell-to-cell adhesion One of the most striking visual phenotypes are the presence of small particles, termed rubies, at the edge of the adherent mucilage both in ruby and mum2 ruby seeds (Figures 3.1 C and D, arrowheads). Based on the shape and size of the particles, it was hypothesised that they were seed coat epidermal cells that had separated from the underlying cell layer (palisade). To test this hypothesis, mature hydrated seeds were stained with the β-glucan stain Calcofluor White in an attempt to observe the seed surface. In wild-type seeds, epidermal cells are in close contact separated only by middle lamellae (Figure 3.3 A), whereas in ruby seeds spaces can be observed between the individual cells and in some places, large cell-sized gaps are apparent on the seed surface (Figure 3.3 B, arrowhead) indicating that cells had detached. The number of the observed gaps correlated with the degree of shaking during hydration (Figure 3.4). To test if the cell-to-cell adhesion also requires Ca2+ bridges formed between HG domains, ruby seeds were treated with a chelator, EDTA, to remove calcium, and CaCl2 to provide it. The addition of EDTA enhanced the loss of epidermal cells (Figure 3.4 F), while the addition of CaCl2 suppressed the loss of epidermal cells (Figure 3.4 E), suggesting that Ca2+ bridges have roles in mediating cell adhesion. Therefore, RUBY activity and calcium bridges both contribute to this process. Since EDTA treatment of wild-type seeds alone does not result in loss of cell-to-cell adhesion (Rautengarten et al., 2008; Western et al., 2001; Voiniciuc et al., 2013), the separation of epidermal cells in ruby suggests that RUBY-mediated cell-to-cell adhesion seems to be more important in the seed coat. 74   Figure 3.3: RUBY is involved in cell-cell adhesion between seed coat epidermal cells as well as between seed coat epidermal cells and palisade cells. (A) and (B) Mature seeds agitated in water for 2 h and stained with Calcofluor White. Bars = 200 μm. (A) Surface of wild-type (Col-2) seed. (B) Surface of ruby-1 seed. White arrowheads indicate spaces where epidermal cells are missing. (C) and (D) Scanning electron micrographs of dry seeds. Bars = 20 μm. (C) Wild type (Col-2), (D) ruby-1.  To determine whether the cell separation occurs before mucilage extrusion, the surfaces of dry wild-type and mutant seeds were examined using scanning electron microscopy. No significant difference in the appearance of middle lamellae of dry mature seeds were apparent between wild type and ruby (n = 6 seeds; Figures 3.3 C and D), indicating that the cell-to-cell adhesion defects are only apparent upon hydration. Indeed, the ruby epidermal cells were observed to lift from the surface of the seed during extrusion (Figure 3.1 O, arrow), 75  corroborating the hypothesis that the cells separate due to mechanical forces generated by the mucilage extrusion.  Figure 3.4 Cell adhesion defects of ruby-1 are enhanced by chelator (EDTA) and suppressed by calcium (CaCl2). ruby-1 seeds gently shaken (A-C) or vigorously mixed on a vortex mixer (D-F), stained by Calcofluor White. (A) Shaken in water. (B) Shaken in 50 mM CaCl2. (C) Shaken in 50 mM EDTA. (D) Vortexed in water. (E) Vortexed in 50 mM CaCl2. (F) Vortexed in 50 mM EDTA.  3.2.4 RUBY is needed to cross-link arabinogalactan-branched RG-I to the seed The visual aspects of ruby phenotype suggest that structural changes occurred in the mucilage and/or middle lamella. To test this possibility, mucilage was extracted from wild-type, ruby, mum2 and mum2 ruby seeds, and the monosaccharide composition determined. Surprisingly, 76  large increases in the amount of rhamnose (Rha; 1.29x), galacturonic acid (GalA; 1.3x), arabinose (Ara; 49.6x) and galactose (Gal; 17x) relative to wild type were observed in the mucilage of ruby seeds (Figure 3.5 A). Stoichiometrically, there was approximately two molecules of Ara and one of Gal for every new molecule of Rha and GalA, suggesting that ruby mucilage contains an arabinogalactan-branched RG-I fraction not previously observed in the wild-type mucilage. This additional pectin may explain why the halo size is increased in the ruby mutant. The appearance of a novel polysaccharide observed in the mutant may have occurred in one of two ways. First, the new RG-I may be present on the surface of wild-type seeds, but not released. Second, the novel RG-I may be synthesised in the mutant, but not the wild type. Third, the branched RG-I may be synthesised in both but processed differently. To distinguish between these hypotheses, a monosaccharide analysis of the whole seed alcohol-insoluble residue (WS-AIR) was completed for wild type and ruby. The levels of Gal, Rha and GalA in wild-type and ruby seeds were similar, suggesting that the ruby mutation results in the release of a branched RG-I that is present but not released from wild-type seed (Figure 3.5 B). The increase in Ara suggests that some of the Ara residues may be absent in the side-chains of the branched RG-I in wild type. To confirm this hypothesis, non-adherent mucilage was extracted with water and then adherent mucilage was extracted from the same seeds with RGase enzyme, and a monosaccharide analysis of these two mucilage layers and the remaining “naked” seeds was performed (Figure 3.5 C). Both mucilage layers were enriched in Ara, Gal, Rha and GalA to the detriment of “naked” seeds in ruby samples, demonstrating that the ruby mutation indeed results in the release of a branched RG-I that is present but not released from wild-type seeds. The modest increase in total Ara in ruby seeds was confirmed. 77   Figure 3.5: Branched RG-I is present in ruby mucilage. (A) Monosaccharide composition of Na2CO3-extracted mucilage as mean values of 4 biological replicates ± SD. (B) Monosaccharide composition of the whole seed alcohol-insoluble residue (WS-AIR) as mean values of 4 biological replicates ± SD. Letters above bars represent groups determined by Tukey’s HSD test, following Welch’s One-way ANOVA performed for each monosaccharide independently. (C) Monosaccharide composition of sequentially-extracted mucilage using water and RGase, and residue left after the extraction. (D) Model of RG-I released in ruby mucilage based on monosaccharide composition, linkage analysis (Table 3.1) and LC-MS/MS analysis of RGase digests (Figure 3.6).  The structure of this novel polysaccharide in ruby mucilage was investigated by carbohydrate permethylated alditol acetate (PMAA) linkage analysis of neutral monosaccharides in the non-adherent mucilage layer. The largest difference between wild type and ruby were in t-78  Araf, 3,6-Gal, and 2,4-Rha (Table 3.1). Their mole percentages and values in nmol mg-1 seed from the monosaccharide composition were used to calculate the ratios to ascertain the structure of the polysaccharide they make. Total mol% values for all PMAAs of a single monosaccharide were added, and percentage of a specific linkage for that given monosaccharide was calculated and multiplied by the absolute values obtained from monosaccharide compositional analysis. For each 2,4-Rha there were 1.1 3,6-Gal, and for each 3,6-Gal there were 2.25 t-Araf, consistent with a branched RG-I where each backbone rhamnose has a molecule of galactose attached to it by a β-(1→4) linkage which in turn has two molecules of t-Araf attached through C-3 and C-6 positions of the pyranose ring. This structure was confirmed by digesting wild-type and ruby adherent mucilage RG-I with RGase and analysing products by high performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD) and Ion Pairing-Reverse Phase (IP-RP)-UHPLC-MS/MS. The digest of wild type showed four distinct peaks with retention time (RT) lower than 15 minutes (Figure 3.6 A, a-d). These peaks were found to contain one to three rhamnoses (R) and two to three galacturonic acids (U), indicating an RG-I with a “naked” backbone with no branching. Additional peaks with higher RT were identified in ruby, but not wild-type mucilage digests (Figure 3.6 B, asterisks). To reveal the structure of these carbohydrate molecules, they were analysed using IP-RP UHPLC-MS/MS. In the first step, the digest of wild type and the digest of ruby were compared by IP-RP-UHPLC-MS (Figure 3.6 C). The main retention process in IP-RP is based on the number of acidic functional groups. In the area of species that contained two acidic functional groups (between RT 8.0 and 11.0 min), a peak, specific to ruby, was identified at RT 9.54 min. This oligosaccharide was selected for IP-RP-UHPLC-MS/MS analysis. Based on the spectrum obtained (Figure 3.6 D) the oligosaccharide that was present in ruby extracts but not in wild-type extracts, which contained 79  the fewest acidic functional groups and the lowest molecular weight, contained two hexoses and four pentoses in addition to two Rha and two GalA residues usually found in digests of RG-I backbone. By using the intense fragment at m/z 747.3, which corresponds to a Z2 and/or B2, it can be postulated that each dimer (Rha-GalA) carries a lateral group consisting of 1 hexose and 2 pentoses. Considering monosaccharide composition and PMAA linkage data, the hexose molecules are most likely galactose, whereas the pentose molecules are most likely arabinose, suggesting an RG-I that has one galactose linked to each rhamnose in β-(1→4) linkage, and two arabinoses linked to each galactose, one in α-(1→3) and one in α-(1→6) linkage (Figure 3.5 D).   80  Table 3.1: Monosaccharide linkage analysis of wild-type and ruby-1 mucilage. The values indicate mean mol% ± SD of two biological replicates. ND, not detected. Major increases are marked in bold. Monosaccharide and Linkage Wild type ruby-1 Fucose        t-Fuc 0.2 ± 0.28 ND Rhamnose        t-Rha 2.15 ± 0.07 0.7 ± 0.28 2-Rha 64.5 ± 1.56 12.2 ± 4.81 2,3-Rha 2.1 ± 0.28 0.5 ± 0.28 2,4-Rha 8.35 ± 4.74 19.15 ± 0.49 2,3,4-Rha 2.45 ± 0.64 1.7 ± 0.14 Arabinose        t-Araf 1.3 ± 1.27 24.25 ± 1.2 t-Arap 0.6 ± 0.14 ND 3-Araf ND 0.3 ± 0 5-Araf 1.55 ± 0.21 1.8 ± 0.57 3,5-Araf ND 1.2 ± 1.41 Xylose        4-Xyl 1.8 ± 0 0.2 ± 0 2,4-Xyl 0.7 ± 0 ND Mannose        t-Man ND 0.1 ± 0.14 2-Man 1.1 ± 0.28 ND 4-Man 1.25 ± 0.35 0.4 ± 0.14 4,6-Man 0.8 ± 0 ND Galactose        t-Gal 1.1 ± 0.14 0.7 ± 0.14 3-Gal 0.55 ± 0.07 1.65 ± 0.07 4-Gal ND 2.9 ± 2.05 6-Gal 0.1 ± 0.14 1.25 ± 0.21 3,6-Gal ND 28.35 ± 4.31 2,4,6-Gal ND 0.3 ± 0.14 3,4,6-Gal ND 0.5 ± 0.14 Glucose        t-Glc 3.25 ± 0.35 0.5 ± 0.28 4-Glc 6.05 ± 0.07 1.65 ± 0.49  81   Figure 3.6: RGase releases novel branched RG-I from ruby-1 mucilage. Mucilage was extracted from wild-type (Col-2) and ruby-1 seeds using RGase enzyme. (A) Chromatogram of wild-type RGase extracts separated using HPAEC. Peaks (a-d) corresponding to non-branched RG-I fragments containing 1 to 3 Rha (R) and 2 to 3 GalA (U) monosaccharides were identified based on standards with identical retention times (data not shown). (B) Chromatogram of ruby-1 RGase extracts. Peaks corresponding to non-branched RG-I fragments 82  are labelled in a-d, and novel peaks are labelled with red asterisks. (C) Base peak ion chromatograms obtained by IP-RP-UHPLC-ESI-MS in negative ionization mode for the digest of wild type (black trace) and the digest of ruby (red trace). (D) IP-RP-UHPLC-ESI-MS/MS spectrum of the oligosaccharide isolated at RT 9.54 as [M-2.H]2- as a precursor ion at m/z 756.23 attributed to the Rha(2)-GalA(2)-Gal(2)-Ara(4) in the digest of ruby mucilage. Red annotations correspond to unambiguous fragments, blue fragments correspond to ambiguous fragments due to the symmetry of the molecule, orange fragments correspond to doubly charged fragments and purple fragments correspond to consecutives fragmentations.  3.2.5 RUBY encodes a putative galactose oxidase The ruby-1 mutation was mapped using a positional cloning approach. 32 mutants were selected from an F2 population made by crossing mum2-1 ruby-1 to wild type Ler, and used to map RUBY to chromosome 1 between DNA markers on BACs F14F17 and F10B61. The genomic DNA of the 32 individuals was also pooled and sequenced, and the low heterozygosity of Col/Ler SNPs used to verify the position on chromosome 1. A mutation in this region was identified in At1g19900. The sequencing of At1g19900 in plants homozygous for two additional alleles of ruby (ruby-2 and ruby-3) also identified mutations suggesting that At1g19900, a gene encoding a putative glyoxal oxidase-related protein, is RUBY (Figure 3.7 A). 83   Figure 3.7: RUBY encodes a glyoxal oxidase-like protein. (A) Schematic of the region of chromosome 1 containing RUBY (At1g19900). Black triangles indicate the position of T-DNA insertions, and arrows show the positions of different EMS-induced point mutations. Numbers under EMS allele labels represent positions of mutations in the coding sequence of RUBY, and letters represent nucleotide changes. Predicted amino acid changes in the deduced protein resulting from the mutation are shown beneath the nucleotide changes. (B) to (E) Ruthenium Red-stained seeds demonstrating that the ruby-5 line with insertion upstream of At1g19900 has a similar phenotype to other ruby alleles. (B) Wild type (Col-0), (C) mum2-10, (D) ruby-5, (E) mum2-10 ruby-5. (F) RT-PCR analysis of RUBY transcript levels in 11 DPA siliques. ACT2 (At3g18780) was used as an internal control. (G) to (I) Ruthenium Red-stained seeds showing that genomic At1g19900 can complement the ruby-1 mutation. (G) mum2-1, (H) mum2-1 ruby-1, (I) ProRUBY:RUBY-Citrine in the mum2-1 ruby-1 background. Bars = 200 μm.   Reverse genetic analysis was used to further verify the identity of RUBY. Seeds from lines homozygous for T-DNA insertions in At1g19900 gene were examined for seed mucilage 84  phenotypes. One line, WiscDsLoxHs097_11H, displayed a phenotype similar to other ruby alleles (Figure 3.7 D, Figures 3.8 A to C). The insertion was confirmed to be 162 bp upstream of the predicted transcription initiation site of At1g19900 (Figure 3.7 A) and designated ruby-5. RT-PCR analysis of At1g19900 using whole siliques at 11 days post anthesis (DPA) suggested that At1g19900 has reduced transcript levels relative to wild type and plants homozygous for other ruby alleles (Figure 3.7 F). The ruby-5 mutation, when introduced into a line carrying a T-DNA allele of mum2 (mum2-10), was able to suppress the mum2 phenotype (Figures 3.7 B to E). An additional line, named ruby-4 (SALK_020627C), showed no phenotype (Figures 3.8 D and E) most likely because it carries an insertion downstream of the At1g19900 coding region (Figure 3.7 A). These data support the hypothesis that At1g19900 is RUBY, and that ruby mutations are able to suppress different alleles of mum2. Additional evidence supporting the hypothesis that RUBY is At1g19900 was obtained by constructing an in-frame fusion of Citrine to the carboxyl terminus of a genomic clone of At1g19900. This genomic clone included the entire 5′ genomic region upstream of At1g19900. When transformed into mum2 ruby plants, this construct successfully complemented ruby, as demonstrated by T2 seeds of all independent transgenic lines exhibiting mum2-like phenotype (Figures 3.7 G, H, and I). 85   Figure 3.8: Insertional mutant, ruby-5, resembles ruby-1 in all phenotypic aspects. Monosaccharide composition of water-extracted mucilage (A) and whole seed alcohol-insoluble residue (B) of wild type (Col-2), ruby-1 and ruby-5. Bars represent means ± SD (n = 3 biological replicates). Letters above bars represent grouping of genotypes based on Tukey’s HSD test (α = 0.05), following Welch’s One-way ANOVA. (C) Comparison of Ruthenium Red-stained mucilage surface of wild type, ruby-1 and ruby-5 seeds (n = 130). Letters represent grouping of genotypes based on Tukey’s HSD test (α = 0.05), following One-way ANOVA (df = 2, F-value = 92.013, p < 0.001). (D) and (E) ruby-4 line does not have ruby-like phenotypes. (D) ruby-4, (E) mum2-1 ruby-4 showing no suppression of mum2.  The identity of RUBY as a glyoxal oxidase was examined by performing a multiple alignment of amino acid sequences and maximum likelihood tree with RUBY and published members of the CAZy (Carbohydrate-Active Enzymes Database) Auxiliary Activity 5 (AA5) family with confirmed activities (Andberg et al., 2017; Aparecido Cordeiro et al., 2010; Avigad 86  et al., 1962; Leuthner et al., 2005; Daou et al., 2016; Kersten, 1990; Kersten and Kirk, 1987; Paukner et al., 2015, 2014; Yin et al., 2015; McPhersons et al., 1992). The phylogram suggests that RUBY forms a clade together with fungal members of the AA5_1 (glyoxal oxidases, EC 1.1.3.15), not the AA5_2 (galactose oxidases, EC 1.1.3.9) subfamily (Figure 3.9 A). Based on amino acids involved in copper coordination and oxidation reaction in the active site, RUBY more closely resembles glyoxal oxidase rather than galactose oxidase (Figure 3.10). A tryptophan residue that was proposed to be necessary for the substrate specificity of GalOx enzymes is missing from the RUBY amino acid sequence, which is replaced by glycine. However, based on research that used site-directed mutagenesis, such a substitution does not completely abolish GalOx activity (Rogers et al., 2007), indicating that it is possible that RUBY is a GalOx.  87   Figure 3.9: RUBY is a putative galactose oxidase functioning on mum2 mucilage. (A) Unrooted Maximum Likelihood tree constructed using amino acid sequence alignment of RUBY with AA5 family enzymes (galactose and glyoxal oxidases) with known functions. Labels at the nodes represent bootstrap support 88  values calculated based on 1000 replicates. Cgl = Colletotrichum gloeosporioides; Cgr, Cg = Colletotrichum graminicola; Fsu = Fusarium subglutinans; Fv = Fusarium verticillioides; Fg = Fusarium graminearum; Fsa = Fusarium sambucinum; Fo = Fusarium oxysporum; Um = Ustilago maydis; Pc = Phanerochaete chrysosporium; Pci = Pycnoporus cinnabarinus  (B) Specific activity of RUBY measured in whole seeds of wild type, ruby-1 and ruby-5 using horseradish peroxidase (HRP) and TMB as a chromogenic substrate for detection of H2O2. Bars represent means (n = 3 biological replicates), error bars represent standard deviations. (C) Oxidation of D-galactose by wild-type seeds assayed using HRP-TMB assay. The control was no treatment prior to activity measurement; Seeds were assayed in the presence of CuSO4, EDTA, Proteinase K, or heat (95°C). Bars represent independently grown biological replicates. (D) Oxidation of Na2CO3-extracted mucilage by commercial galactose oxidase. H2O2 was measured using HRP-TMB. The mucilage samples were reduced using NaBH4 (reduced) or left untreated (control). Bars represent means ± SD of 3 independently grown biological replicates. Letters above bars represent groups based on Tukey’s HSD test (α = 0.05), following Two-way ANOVA. The main effect of genotype on substrate availability was significant (df = 3, F-value = 153.670, p < 0.001), as well as the effect of treatment (reduction) on the substrate availability (df = 1, F-value = 13.347, p < 0.005), and interaction between genotype and treatment (df = 3, F-value = 12.966, p < 0.001). (E) to (H) Extracted chromatograms obtained by IP-RP-UHPLC-MS for the four genotypes (wild type [Col-2], black trace; mum2-1, red trace; ruby-1, blue trace; mum2-1 ruby-1, green trace). Oligosaccharides presented are labelled with regards to the number of R (Rha), U (GalA), and G (Gal). (E) R3U3 isolated as [M-H]- at m/z 983.23; (F) R3U3G3 isolated as [M-2H]2- at m/z 734.18; (G) R3U3G1 isolated as [M-H]- at m/z 1145.29 and (H) oxidised R3U3G1 isolated as [M-H]- at m/z 1143.27. Exact masses of each compound were selected with a mass window of ± 0.1 Da.   89  Table 3.2: A list of compounds tested as substrates for oxidases on dry mature seeds using HRP-TMB assay. + positive reaction; - no reaction; * weak reaction Compound Activity Wild type ruby-1 D-Galactose + - D-Galacturonic acid + + D-Glucuronic acid + + Glycerol + - Meso-erythritol + - o-Nitrophenyl β-D-Galactopyranoside + - p-Nitrophenyl β-D-Galactopyranoside + - Raffinose + - Lactose* + - Butan-1-ol - - D-Glucose - - D-Mannose - - D-Xylose - - Ethanol - - Galactan - - Galactomannan - - Glyoxal - - Glyoxylic acid - - Guar - - Isopropanol - - L-Arabinose - - L-Fucose - - L-Rhamnose - - Mannitol - - Methanol - - Methylglyoxal - - Myo-inositol - - Sorbitol - - Sucrose - - Trehalose - - Xyloglucan - -   90  To test the enzymatic activity of RUBY, it was expressed in E. coli. Protein fused with C-terminal His6 tag was partially soluble when expressed at 11°C in Arctic Express (DE3) cells (Figure 3.11), but the ensuing protein showed no activity against monosaccharides, glyoxal, methylglyoxal or glyoxylic acid. However, dry wild-type seeds were able to generate hydrogen peroxide (H2O2) in the presence of glycerol, whereas ruby-1 and ruby-5 seeds showed no such activity. Multiple monosaccharides, disaccharides, trisaccharides, polyols, alcohols and carbonyl compounds (Table 3.2) were tested as substrates, and the reaction was further investigated for the compounds showing positive reaction. The only monosaccharide oxidised by RUBY was galactose, suggesting that RUBY is a GalOx (Figure 3.9 B). The reaction was inhibited by heating at 95°C and by Proteinase K, suggesting that the reaction is dependent on a protein (Figure 3.9 C). Pre-incubation in CuSO4 substantially increased the activity, as has been previously observed for fungal GalOxs (Spadiut et al., 2010), and also reduced differences between biological replicates (Figure 3.9 C). This suggests that RUBY is activated by Cu2+, a co-factor in the active sites of GalOx enzymes. Likewise, pre-incubation in EDTA, a chelator of divalent cations, reduced the activity by half, but the difference is not statistically significant (Figure 3.9 C). Like fungal GalOx enzymes, RUBY uses glycerol, and also meso-erythritol, as a substrate (Figure 3.9 B). In addition, galactose-containing raffinose was oxidised, consistent with previous reports of GalOx enzyme activity (Paukner et al., 2015, 2014; Avigad et al., 1962). Oxidation of lactose was observed, but the reaction was very weak, again consistent with other GalOx enzymes (Xu et al., 2000; Avigad et al., 1962). For this reason, the analysis of the specific activity of the enzyme on lactose was not feasible. To analyse if RUBY has a preference for the Gal α-anomer over the β-anomer, we tested activity on p-nitrophenyl-β-D-Gal (PNP-β-D-Gal) and o-nitrophenyl-β-D-Gal (ONP-β-D-Gal). Both were readily used by RUBY (Figure 3.9 B), 91  suggesting that RUBY can use Gal regardless of anomerism. Galactose-containing polysaccharides – galactomannan, guar gum, xyloglucan and linear galactan were not oxidised by RUBY. Since the assays were done on seeds, it is possible that these large substrates could not access the enzyme, which is likely inside the cellulosic secondary cell wall (see below).   92    Figure 3.10: RUBY contains catalytic amino acids required for galactose/glyoxal oxidase function. Multiple alignment of amino acid sequences visualised by JalView using BLOSUM62 colouring scheme for indicating conserved amino acids. Catalytic amino acids necessary for glyoxal oxidase and galactose oxidase function are indicated by red arrowheads. Tryptophan found in the active site of galactose oxidases, but not glyoxal oxidases is indicated by a black arrowhead. 93    Figure 3.11: Western blot analysis of purification of RUBY-His6 expressed in Arctic Express (DE3) strain of E. coli. Proteins were extracted from cells and purified on Ni-NTA column. Fractions were analysed using Western blot. (A) Ponceau S-stained nitrocellulose membrane. (B) His-Probe (H3) labelling of proteins. Numbers designate lanes containing: supernatant after lysis (1), pellet after lysis (2), molecular weight marker (3), column flow-through after binding (4), wash 1 (5), wash 2 (6), wash 3 (7), wash 4 (8), wash 5 (9), wash 6 (10), wash 7 (11), eluate 1 (12), eluate 2 (13), eluate 3 (14), eluate 4 (15).  Interestingly, apart from having more Gal than wild type, mum2 mucilage was found to have oxidised Gal residues attached to Rha (Macquet et al., 2007b). Mucilage was extracted with Na2CO3 and tested for the amounts of oxidisable Gal residues using commercial GalOx. As expected, mum2-1 and mum2-1 ruby-1 have the highest amounts of oxidisable Gal (Figure 3.9 D) due to an increase in t-Gal. This indicates that RUBY can oxidise mum2 mucilage, but it may not act on wild-type mucilage, as suggested by the low levels of oxidation and lack of difference 94  between wild type and ruby-1. To determine if RUBY functions to oxidise mum2 mucilage to make it insoluble, extracted mucilage samples were divided into two, putative aldehydes reduced in one sample using NaBH4, and the other sample processed as a control without reduction. Strikingly, reduced mum2 mucilage showed two times more available substrate than control (Figure 3.9 D), suggesting that approximately half of the Gal was oxidised in extracted mum2 mucilage. This increase was not observed in reduced mum2 ruby mucilage compared to control, suggesting that the oxidation of mum2 mucilage is dependent on RUBY.  Adherent mucilage from wild type, ruby-1, mum2-1, and mum2-1 ruby-1 was analysed further. Non-adherent mucilage was first extracted using mild acid and mild alkali sequentially, and further hydrolysed the adherent mucilage surrounding seeds with rhamnogalacturonan hydrolase, as described by Macquet et al., (2007b). Hydrolysates were analysed on IP-RP-UHPLC-MS. Besides unbranched RG-I oligosaccharides (mainly R2U2 and R3U3 that were present in all samples; Figure 3.9 E), several galactosylated RG-I oligosaccharides were detected in mum2-1 and mum2-1 ruby-1 (Figures 3.9 F and G), R3U3G1 being particularly abundant (Figure 3.9 G). Interestingly, R3U3G1 is present in both mum2-1 and mum2-1 ruby-1 (Figures 3.9 G), but oxidised forms of this oligosaccharide are present in mum2-1 only (Figure 3.9 H). This shows that some Gal units are indeed oxidised in mum2 mucilage, and that this oxidation of mum2 mucilage is dependent on RUBY.  3.2.6 RUBY is expressed in seeds after mucilage secretion and localises to the apoplast in columella The ProRUBY:RUBY-Citrine construct that complemented ruby was used to study the temporal and spatial expression, and the subcellular localisation of RUBY. Developing seeds of T2 plants 95  were removed from siliques and imaged by spinning-disc confocal microscopy. No signal was observed before 9-10 DPA, indicating that RUBY is expressed following the completion of mucilage secretion. The signal at 10 DPA was localised in the secondary wall of developing columella, and in the primary cell walls and middle lamellae around the cells (Figure 3.12 A). At 13 DPA when the columella is fully developed (Western et al., 2000), signal continues to accumulate in the columella and primary cell walls surrounding epidermal cells (Figure 3.12 B). The signal does not lose intensity even in fully developed dry seeds. RT-PCR analysis also demonstrated that RUBY is primarily expressed in siliques late in the development (Figure 3.13 D). These results demonstrate that RUBY, consistent with its roles in mum2 mucilage cross-linking and cell-to-cell adhesion, localises to the columella adjacent to the mucilage pocket, as well as to the primary cell walls surrounding cells. The signal is also visible around the cells in the underlying palisade cell layer (Figure 3.12 B), and in the epidermis of the mature root (Figures 3.13 B and C). However, the fluorescence in the root was weak, consistent with no detected transcript in this tissue (Figure 3.13 D). 96   Figure 3.12: RUBY-Citrine is expressed after mucilage secretion and localises to the apoplast. Developing seeds carrying ProRUBY:RUBY-Citrine in mum2-1 ruby-1  background imaged on a spinning disc confocal microscope. (A) Seed surface at 10 DPA. (B) Seed surface at 13 DPA. Bars = 20 μm.  (C) to (E) Top view of the seed surface stained with FM4-64 dye. Bars = 5 μm. (C) RUBY-Citrine shown in yellow. (D) FM4-64 imaged shown in magenta. (E) Overlay of RUBY-Citrine (yellow) and FM4-64 (magenta) demonstrating that RUBY-Citrine localises outside of the plasma membrane. c, columella; m, mucilage pocket; arrowhead, middle lamella.  To test whether RUBY is indeed secreted, we examined localisation in the apoplast by staining plasma membrane of ProRUBY:RUBY-Citrine developing seeds with the dye FM4-64. In the overlay of RUBY-Citrine (yellow; Figure 3.12 C) and plasma membrane (magenta; Figure 97  3.12 D) images, it is evident that RUBY localises outside of the plasma membrane (Figure 3.12 E), demonstrating extracellular localisation of the protein. At earlier stages of development (9-10 DPA), it is possible to observe fluorescence inside the cells in punctate or reticulate patterns, most likely representing the protein in the secretory pathway prior to secretion.  Figure 3.13: RUBY is expressed in root epidermis and seed coat. Confocal microscopy images of ProRUBY:RUBY-Citrine root. Images represent parts of a single root: (A) root tip, (B) section between elongation and maturation zone (appearance of root hairs), (C) mature root. Bar = 100 μm. (D) RT-PCR showing tissue-wide expression of RUBY. GAPC1 (At3g04120) was used as an internal control.   98  3.2.7 Pectin cross-linking is most likely mediated through hemiacetals, not through hydroxycinnamate esters Cross-linking of RG-I via dimerisation of ferulic acid (FA) attached to arabinose and galactose side-chains of sugar beet RG-I, or arabinose side-chains of arabinoxylan, has been previously reported (Saulnier and Thibault, 1999; Ralet et al., 2005; Grabber et al., 1995; Fry, 2004). Oxidative coupling of arabinoxylan-FA occurs in the presence of H2O2 and peroxidases (Encina and Fry, 2005; Burr and Fry, 2009). Since the RUBY reaction generates H2O2, we investigated whether RUBY functions to cross-link cell walls through dimerisation of hydroxycinnamate esters.   Mature seeds were treated with 2 M NaOH to extract any ester-linked phenolic compounds that are present in the mucilage and the columella surface, and analysed extracts by HPLC-UV.  The most abundant phenolic compound detected was sinapic acid, confirmed by comparison to retention time (Figure 3.14 A) and UV absorbance of a standard, as well as a molecular mass [M-H]- of 223.0599 (Figure 3.15 C). Like FA, sinapic acid can also form dimers (Bunzel et al., 2003). If wild type, expressing functional RUBY, makes dimers of sinapic acid, it would be expected to have lower levels of sinapic acid (monomers) than ruby. However, we did not observe differences in sinapic acid between wild type and ruby (Figure 3.15 A), suggesting that it exists only as a monomer on the seed surface. A second compound was detected and was reduced by ~30% in ruby compared to wild type (Figure 3.15 B). Based on a molecular mass [M-H]- of 447.0906 (Figure 3.15 C) and published results on Arabidopsis seed phenolics (Routaboul et al., 2006), this compound is most likely quercetin-3-O-rhamnoside (Q3R).   99   Figure 3.14: RUBY most likely functions through hemiacetal formation, not through hydroxycinnamate cross-linking. (A) HPLC-UV chromatograms representing sinapic acid standard (black) and surface phenolics of wild type (Col-2) seeds released by 2 M NaOH (blue). Vanillin was added as an internal standard. (B) Quantification of sinapic acid released from seed surface of mutants for genes in the sinapic acid biosynthetic pathway. Bars represent mean ± SD of 3 independently grown biological replicates. Letters 100  above bars represent groups assigned based on Tukey’s HSD test, following One-way ANOVA (df = 8, F-value = 21.939, p < 0.001). (C) to (F) Seeds agitated in water and stained with Ruthenium Red. (C) Col-0 (wild type), (D) fah1-2 mutant (Col-0 background), (E) Ler (Wild type), (F) fah1-7 mutant (Ler background). (G) Photograph of re-hydrated mucilage samples in 96-well plate after Na2CO3 extraction, reduction, and purification, demonstrating solubility/insolubility of mucilage. (H) Na2CO3 can break cross-links made by RUBY. Mucilage extracted from mum2-1 seeds with Na2CO3, purified, dried and re-hydrated. One sample was mixed with water (Control) and the other with Na2CO3. (I) to (L) Reduction promotes mucilage extrusion from mum2 seeds. Seeds were incubated with NaBH4 at neutral pH. (I) Wild type (Col-2) without NaBH4, (J) Wild type (Col-2) with NaBH4, (K) mum2-1 without NaBH4, (L) mum2-1 with NaBH4. (M) to (P) Drying promotes insolubility of mum2 mucilage through aldehydes. Images show re-hydrated seeds after base-extraction of mucilage, reduction, and air-drying. (M) Wild type (Col-2) without NaBH4, (N) Wild type (Col-2) with NaBH4, (O) mum2-1 without NaBH4, (P) mum2-1 with NaBH4. Scale bars = 200 μm.   To test if lack of sinapate has an effect on the whole seed mucilage phenotype, we stained seeds of mutants known to be involved in sinapic acid biosynthesis, with Ruthenium Red. Based on HPLC-UV quantification, fah1-2 and fah1-7 mutants have almost complete reduction in sinapic acid (Figure 3.14 B). However, we observed no difference in seed mucilage phenotype between these mutants and wild type (Figures 3.14 C to F), indicating that hydroxycinnamates are likely not involved in RUBY-mediated cross-linking.  Further investigation of mechanisms by which RUBY may function was guided by the observation that purified non-reduced (control) mum2 mucilage was unable to fully hydrate in water, resulting in increased opacity of the solution (Figure 3.14 G). The reduced mum2 sample, however, re-hydrated to a higher degree, resulting in a transparent solution (Figure 3.14 G). This result suggests that oxidation reduces the solubility of mum2 mucilage. When mixed with Na2CO3, mum2 mucilage became more soluble, as evident by a loss of opacity (Figure 3.14 H). 101  This result demonstrates that the cross-links making oxidised mucilage hydrate poorly can be disrupted by Na2CO3. To determine the relevance of Gal oxidation for mucilage extrusion, we tested whether the reduction of carbonyls by NaBH4 can release mucilage from mum2. NaBH4 reductions are usually performed in basic solutions to prevent its decomposition, but basic solutions can also extract mucilage from mum2 seeds, which would obscure the effects of reduction on mucilage hydration properties. To avoid using basic solutions and to prevent pH shift upon addition of NaBH4, we performed reduction in imidazole-HCl buffer at pH 7, which was previously suggested to increase stability of NaBH4 (Kim and Carpita, 1992). We observed a release of mucilage from mum2-1 seeds in the presence of the reductant (Figure 3.14 L), whereas control seeds showed patchy extrusion only sporadically (Figure 3.14 K). The wild-type seeds displayed no clear difference with regards to the treatment (Figures 3.14 I and J). This result indicates that the oxidation of Gal into an aldehyde makes mucilage insoluble in vivo. It has been suggested that polysaccharides oxidised by GalOx can form insoluble aerogels when dried through the formation of hemiacetals (Mikkonen et al., 2014). To test if drying can make the mucilage insoluble, we reduced wild-type and mum2 seeds in a basic solution to ensure mucilage extrusion, air-dried them, and re-hydrated them in water. The mucilage of mutant seeds re-hydrated only after reduction with NaBH4 (Figure 3.14 P), whereas mucilage of non-reduced control remained collapsed (Figure 3.14 O). Similar to other experiments, wild type showed no difference between treatments (Figures 3.14 M and N). The fact that oxidised mucilage becomes insoluble when dried, suggests that hemiacetal formation may be responsible, as has been observed in studies of aerogels. The mucilage of mum2 has more RG-I Gal side-chains than that of wild type. As a consequence, RUBY has more substrate 102  available, which, once oxidised, can react with hydroxy groups of highly abundant polysaccharides in their environment (RG-I) to form hemiacetals (Figure 3.16).   Figure 3.15: Most abundant phenolics on seed surface are sinapic acid and a flavonol glycoside. (A) to (B) UV detector response as peak intensity measurements of two most abundant compounds released from the seed surface by saponification in NaOH. Values were adjusted for losses using internal standard (vanillin). (A) Sinapic acid peak intensities. Bars represent mean ± SD of 3 independently grown biological replicates. Letters above bars represent groups assigned based on Tukey’s HSD test, following One-way ANOVA (df = 3, F-value = 5.6113, p < 0.05). (B) Flavonol glycoside peak intensities. Bars represent mean ± SD of 3 independently grown biological replicates. Letters above bars represent groups assigned based on Tukey’s HSD test, following One-way ANOVA (df = 3, F-value = 21.703, p < 0.001). (C) to (D) Analysis of seed surface phenolics using LC-ESI-MS in negative mode. Mass spectra represent Peak1 (B) and Peak 2 (C) from the chromatogram shown in Figure 3.14 A.  103   Figure 3.16: A model proposing cross-linking of branched RG-I by galactose oxidation. (A) In the seed coat of drying seeds RUBY is distributed in the apoplast surrounding epidermal cells. Based on results of experiments performed on mum2 mucilage, we propose that RUBY oxidises terminal Gal to generate carbonyl group that reacts with hydroxy groups of polysaccharides in its proximity (e.g. unbranched RG-I) to create hemiacetal bonds (B). The proposed substrate in the middle lamella is branched RG-I with single Gal side-chains, which lacks t-Ara residues in the presence of RUBY.   104  3.3 Discussion We have identified a gene, RUBY, encoding a protein associated with galactose oxidase activity that is required for normal cell-to-cell adhesion and mucilage structure in the seed coat of Arabidopsis. Our data suggest that RUBY promotes cross-linking of RG-I in the middle lamella and mucilage via oxidation of galactose and, presumably, subsequent formation of a hemiacetal between oxidised Gal and polysaccharides in its proximity. Taken together our results define a role for galactose oxidases in the cross-linking of cell wall carbohydrates for the purposes of strengthening cell-to-cell adhesion and possibly mucilage cohesion.  3.3.1 RUBY is a putative galactose oxidase with RG-I side-chains as its substrate in the mucilage The Arabidopsis protein RUBY has an amino acid sequence similar to a family of fungal oxidases that use small carbonyl compounds or galactose as substrates. RUBY is secreted by seed coat epidermal cells late in seed development and remains there throughout maturation. Intact mature seeds of wild type, but not ruby mutants, display galactose oxidase activity. These data suggest that RUBY is likely a galactose oxidase that is secreted by seed coat epidermal cells late in development. Fungal galactose oxidases have been well-characterized enzymatically. Such proteins oxidise the hydroxy moiety at the C-6 carbon of galactose, either as a monosaccharide or a terminal component of a galactose-containing polysaccharide, to produce an aldehyde and H2O2 as a by-product (Avigad et al., 1962). However, the biological function of fungal galactose oxidases is unknown. Here, we showed that the plant galactose oxidases strengthen the middle lamella and seed mucilage, supporting a role in the cross-linking of cell wall carbohydrates.  105  Although the exact substrate of RUBY in wild-type seed coat epidermal cells is not known, several lines of evidence suggest that RUBY oxidises galactose in RG-I side chains. First, RUBY promotes cell-to-cell adhesion, which normally occurs via the pectin of the middle lamella, and RG-I is a pectin with an abundance of galactose side-chains. Second, mutations in RUBY suppress the mucilage extrusion phenotypes of mum2 and bxl1. Both MUM2 and BXL1 encode exoglycosidases that hydrolyse the side chains of RG-I. Third, mutations in ruby result in the release of a branched RG-I from the seed coat epidermal cells. This RG-I is normally present in wild-type seeds, but not released, suggesting that RUBY functions to cross-link the RG-I to the cell surface. Finally, RG-I in mum2 mucilage contains oxidised galactoses whose formation is dependent on RUBY (Macquet et al., 2007b; Figures 3.9 D, G and H).  The branched RG-I extracted with the mucilage of ruby mutants has a distinct structure, not previously described, where each molecule of rhamnose is covalently bonded to one molecule of β-D-Gal which in turn is linked, via carbons 3 and 6, to two molecules of Ara (Figure 3.5 D). This RG-I appears to be present in wild-type epidermal cells, but is extracted from the surface of the seed with the mucilage only in the absence of functional RUBY. These data suggest that RUBY cross-links branched RG-I to the seed coat epidermal cell even though the molecule has no terminal Gal substrate to be oxidised. This cross-linking could be explained in one of two ways. The terminal Ara residues on the RG-I side chains might form hemiacetals with oxidised Gal on other carbohydrates. Alternatively, we did observe a significant increase in total Ara in ruby versus wild-type seeds (Figure 3.5 C), suggesting that the Ara is added to the branched RG-I more extensively in the ruby mutant. If so, the branched RG-I present in wild type that is bound to the seed surface may lack many of the terminal Ara molecules observed in 106  ruby mucilage and instead have primarily Gal side-chains, making it a possible direct substrate for RUBY.   3.3.2 Oxidation of galactose most likely creates hemiacetal bonds in the mucilage The oxidation of galactose could promote carbohydrate cross-linking through at least two non-mutually exclusive mechanisms. First, it has been shown in a variety of plant species that hydroxycinnamic acids can be covalently bonded to Gal or Ara and then oxidatively cross-linked by H2O2 and peroxidases (Bunzel et al., 2003; Encina and Fry, 2005; Fry, 2004; Grabber et al., 1995; Ralet et al., 2005; Ralph et al., 1994; Saulnier and Thibault, 1999; Burr and Fry, 2009). Therefore, the availability of Gal or Ara sidechains on RG-I in the apoplast of seed coat epidermal cells could provide substrate for the covalent bonding to a hydroxycinnamic acid. Oxidation of galactose by RUBY would generate the H2O2 needed to cross-link two molecules of hydroxycinnamic acid attached to different carbohydrate chains. While this hypothesis is consistent with much of our data, we were unsuccessful in our attempt to find direct evidence to support the involvement of hydroxycinnamic acids as a structural element in either middle lamellae or mucilage. Sinapic acid was the only hydroxycinnamic acid we could detect in seeds, and fah1-2 and fah1-7 mutant seeds lacking sinapic acid (Figures 3.14 C to F) did not show seed coat epidermal defects in either cell-to-cell adhesion or mucilage cohesion. These data suggest that cross-linking of hydroxycinnamic acids is not a mechanism used in the apoplast of seed coat epidermal cells. Like hydroxycinnamic acids, tyrosine amino acids present in HRGPs could be cross-linked in the presence of H2O2 and peroxidases (Fry, 2004; Waffenschmidt et al., 1993; Kjellbom et al., 1997). Thus, it is also possible that the H2O2 generated by RUBY is used to crosslink HRGPs present in the mucilage and middle lamellae. HRGPs have been identified in 107  seed mucilage through proteomic analyses, but mutations in genes encoding such proteins exhibit no phenotypes (Tsai et al., 2017). A second possibility for the formation of cross-links through galactose oxidation is the formation of hemiacetals between oxidized galactose and hydroxy groups on neighbouring polysaccharides (Merlini et al., 2015; Parikka et al., 2012). Such direct cross-linking of polysaccharides in the plant cell walls has not yet been shown, but at least two studies have suggested that enzymatic oxidations of hemicellulosic polysaccharides in vitro can lead to such cross-linking. Both galactomannan (GM) and xyloglucan (XyG) formed gels in the presence of GalOx, horseradish peroxidase (HRP) and catalase (Parikka et al., 2010), whereas fenugreek GM formed a gel when treated with combination of laccase and TEMPO (2,2,6,6-tetramethyl-1-piperidinyloxy radical) (Rossi et al., 2016). In both cases Gal side-chains were oxidised at the C-6 position to aldehydes, and hemiacetal formation with another hydroxy group in the proximity was proposed as a cross-linking mechanism. 2D-NMR spectroscopy showed the existence of the hemiacetal bond between oxidised galactose and C-4 of mannose in the backbone of fenugreek GM (Merlini et al., 2015). These data indicate that Gal oxidation could directly result in the formation of stable hemiacetal cross-links and, therefore, that RUBY could generate cross-links in the apoplast in the presence of pectin-rich environments like mucilage and the middle lamellae. Hemiacetal formation and breakdown are both catalysed by acids and bases (Ernst and Inge, 1967). Our observation that a base, Na2CO3, can disrupt cross-linking in the mum2 mucilage (Figure 3.14 H) suggests that the carbonate extraction of mum2 mucilage may be due to hemiacetal breakdown. This hypothesis is further supported by the fact that NaBH4, a reducing agent for carbonyls that can reduce Gal aldehydes to prevent formation of hemiacetals, increases solubility of mum2 mucilage (Figures 3.14 K and L). Additionally, oxidation of monosaccharides 108  into aldehydes in polysaccharides results in the formation of insoluble aerogels upon drying, owing to hemiacetal formation (Christensen et al., 2001; Ghafar et al., 2015; Köhnke et al., 2014; Mikkonen et al., 2014). Our data suggest that once the base is removed by washing or dialysis and samples dried, cross-links are re-formed and mucilage becomes less soluble again (Figures 3.14 G, O and P). This observed reduced solubility of mum2 mucilage can be prevented by the reduction of aldehydes (Figures 3.14 O and P). Therefore, we propose that RUBY oxidises terminal Gal residues on RG-I to create aldehydes. In the cell wall, an environment rich in carbohydrates, the abundance of hydroxy groups around newly formed aldehydes may promote formation of hemiacetals. Once the pectin is extracted with basic solutions, it is necessary to dry it to bring these two functional groups together and re-form hemiacetals. Cross-linking through hemiacetals would have an advantage over hydroxycinnamate or HRGP cross-linking because it requires only a single enzyme. RUBY functions at the time when the epidermal cells are undergoing programmed cell death, thus the generation of the H2O2 as a by-product may not be harmful to the seed, eliminating the need for peroxidases.  Creating hemiacetals by GalOx enzymes can be beneficial in tissues where the cell wall needs additional reinforcements due to a high exposure to mechanical stress. In the seed coat epidermis, the middle lamella must resist the shear forces generated by rapid extrusion of the mucilage. Indeed, we have shown that one role of RUBY is to strengthen the middle lamellae between adjacent seed coat epidermal columellae, as well as between seed coat epidermal cells and the underlying palisade. The fact that, in ruby mutants, cell separation was not evident in mature dry seed (Figures 3.3 C and D), but was obvious only following hydration and mucilage extrusion (Figures 3.3 A and B) suggests that ruby cell separation requires mucilage extrusion.  109  3.3.3 Mutations in RUBY affect mucilage appearance In addition to the attachment of the columellae to the seed surface, RUBY appears to be required for the connection of the primary cell wall to the top of the columellae. In wild-type cells mucilage extrusion breaks the radial portion of the primary wall. The resulting cell wall fragment remains firmly attached to the top of the columellae (Western et al., 2000; Figure 3.1 N). In contrast, these cell wall fragments typically separate from the columellae and are observed within the adherent mucilage of ruby mutants (Figure 3.1 N), suggesting that RUBY strengthens connections between the primary wall and the columellae. This hypothesis is consistent with the appearance of RUBY in the columellae late in seed coat differentiation (Figures 3.12 A and B). In contrast to the middle lamellae, mucilage needs to expand upon hydration, so cross-links throughout mucilage would be detrimental to its ability to extrude during hydration. Our data suggest that the removal of RG-I galactose side chains by MUM2 is necessary to allow mucilage extrusion in the presence of active RUBY. However, it is less clear whether RUBY influences wild-type mucilage pectin once MUM2 has removed the galactose side chains. The mucilage of the ruby single mutant has a dishevelled appearance and a larger adherent mucilage halo (Figure 3.1 P; Figure 3.8 C), indicating that the mucilage is not normal. It is possible that the removal of galactose side-chains from RG-I by MUM2 is not complete, and that RUBY protein in the columellae adjacent to the mucilage pocket (Figures 3.12 A and B) establishes the correct cohesiveness of wild-type mucilage. However, the dishevelled mucilage could be an indirect effect of the loosened epidermal cells and/or primary cell walls detaching from the columella, whereas the larger halo may be the result of the release of additional pectin from the middle lamellae. 110  One curious aspect of the ruby mutant phenotype is the collapsed cellulosic rays observed in the adherent mucilage (Figures 3.1 Q and R). Collapsed rays have been observed in mutants that fail to synthesise the mucilage galactoglucomannans (GGM) that surround the cellulose rays of extruded mucilage (Yu et al., 2014; Voiniciuc et al., 2015b). Since the galactose side-chains of GGM are a potential substrate of RUBY, it is tempting to speculate that RUBY may play a role in the strengthening of the mucilage ray structure, but confirmation of such a role would require additional evidence.    In addition to the seed coat epidermal cells, RUBY is expressed at low levels in the root epidermis starting at the elongation zone (Figures 3.13 A to D). To date we have been unable to identify ruby phenotypes in these cell types. RUBY is one of seven homologous genes present in the Arabidopsis genome, so it is possible that redundancy obscures some mutant phenotypes. Molecular genetic and biochemical analyses of these RUBY homologs may shed additional light on the role of galactose oxidation in pectin cross-linking, as discussed in the following chapter.  111  Chapter 4: Functional analysis of the GALACTOSE OXIDASE-LIKE (GOXL) gene family from Arabidopsis thaliana 4.1 Introduction Our research has indicated that the RUBY (At1g19900), a putative galactose oxidase, plays a role in forming covalent bonds between galactose and monosaccharides on adjacent carbohydrates. There are six additional genes in the Arabidopsis thaliana genome annotated as encoding glyoxal-oxidase-related proteins. Studying these genes should expand our knowledge of both the biochemistry and biological roles of these enzymes. In this chapter, I explore the expression, and biochemical and biological function of these six RUBY homologues.  Only one of the RUBY homologues, GLOX1/GOXL1 (At1g16729), was studied prior to RUBY. GOXL1 was shown to be a direct target of the MYB80 transcription factor, which is proposed to repress GOXL1 expression prior to stage 8 or 9 of anther development (Phan et al., 2011). GOXL1 expression is most prominent in later stages of anther development and in pollen grains, but no genetic or biochemical studies were done to understand its function. Since, during my thesis research, I found that GOXL1 is involved in pollen function, below I review pollen development and the pollen wall composition and structure.  4.1.1 Pollen development In Arabidopsis, pollen development occurs inside the locules of anthers. Each anther, which sits on top of a filament in a fully developed stamen, consists of four locules (Figure 4.1). Pollen development begins with meiosis, resulting in tetrads of haploid microspores (Figure 4.1, Td) each of which develops into a pollen grain. At stage 8 of anther development, which corresponds 112  to the stage 10 of flower development (Smyth et al., 1990), microspores are released from tetrads and pollen grain development begins (Sanders et al., 1999). This separation requires removal of a thick callose-rich wall that has been deposited by the microspore mother cell as well as the microspores themselves (Owen and Makaroff, 1995). During stage 9, the pollen wall exine forms (Sanders et al., 1999). The substrate for the exine is provided exogenously by the tapetum, a nutritive cell layer, which lines the inside of locules (Figure 4.1, T; Chapman, 1987). At the stages 10 and 11, which are close to stages 11 and 12 in flower development, the tapetum undergoes programmed cell death, and the stomium, a specialised structure made from epidermal cells that is important for anther opening during dehiscence (Wilson et al., 2011; Sanders et al., 1999), starts to form (Sanders et al., 1999). At stage 11, the microspore undergoes an asymmetric cell division (Pollen Mitosis I) to give a generative and a vegetative cell (Owen and Makaroff, 1995). This is followed by Pollen Mitosis II, where the generative cell divides once more to give a mature, tricellular, pollen grain (Borg et al., 2009). At stage 12 of anther development, corresponding to stage 12 of floral development, the pollen grains are developed, the stomium is differentiated and the septum between the locules breaks, resulting in the fusion of adjacent locules (Figure 4.1, Stage 12; Sanders et al., 1999). At this point the development is completed, and subsequent stages encompass the dehydration of pollen grains and anther dehiscence, resulting in the release of pollen. 113   Figure 4.1: Schematic representation of transverse section through an anther during pollen development. C, connective; E, epidermis; En, endothecium; ML, middle layer; S, septum; St, stomium; StR, stomium region; T, tapetum; Td, tetrads; TPG, tricellular pollen grains; V, vascular bundle. Reprinted with permission. Scott, R.J., Spielman, M., and Dickinson, H.G. (2004). Stamen Structure and Function. Plant Cell 16: 46–60. www.plantcell.org. Copyright American Society of Plant Biologists.   114  4.1.2 Pollen wall structure and composition One of the most prominent events during the pollen development is the deposition of the pollen wall. Following meiosis of the pollen mother cell into tetrads, microspores deposit a cellulose-rich primexine layer underneath the callose wall (Figure 4.2; Heslop-Harrison, 1968; Shi et al., 2015). Primexine serves as a receptor for sporopollenin, a material used for exine synthesis (Shi et al., 2015). Recent studies have demonstrated that primexine contains AGPs and xylans, both of which are required for proper sporopollenin deposition and exine architecture (Li et al., 2017; Suzuki et al., 2017). As the callose is degraded, miscrospores separate and expand due to the lack of constraint from the callose (Figure 4.2). This results in thinning and shredding of the primexine, followed by the synthesis of new layers of pollen wall (Heslop-Harrison, 1968a).  Figure 4.2: Development of the pollen wall. Figure based on Ariizumi and Toriyama, (2011) and Shi et al., (2015).  After callose is removed, exine and intine are synthesised (Figure 4.2). Exine is the outermost layer of the pollen wall, which is made of two distinct layers – outer, sexine, and inner, nexine (Figure 4.3; Heslop-Harrison, 1968a). In Arabidopsis, the sexine has characteristic reticulate pattern, resulting from localised deposition of sporopollenin into structures containing 115  rod-like bacula with tectum on top (Figure 4.3; Paxson-Sowders et al., 1997). The chemical composition of sporopollenin is not fully understood due to its resistance to chemical and physical degradation, but several genetic and biochemical studies done on Arabidopsis indicate that it contains hydroxylated polyketides and hydroxycinnamoyl spermidines (Kim et al., 2010; Grienenberger et al., 2010, 2009; Quilichini et al., 2014). The space between the baculae of mature pollen contains tryphine or pollen coat (Figure 4.3). It is composed of saturated fatty acids, alkanes, flavonoids, carotenoids and proteins (Hsieh and Huang, 2007; Piffanelli et al., 1998) that are released upon tapetal degradation, and has roles in adhesion of the pollen grain onto surfaces, including the stigma (Piffanelli et al., 1998). At the bottom of the baculae is the foot layer, also known as nexine I, which can be distinguished from nexine II using transmission electron microscopy (TEM; Heslop-Harrison, 1968a). Nexine I is deposited by the tapetum between baculae, whereas nexine II is synthesised by the microspore itself (Heslop-Harrison, 1968a). Similar to exine, nexine contains sporopollenin, but it also cannot form without AGPs  (Jia et al., 2015). A layer underneath the nexine is the intine (Figure 4.3), which is fully gametophytic in origin (Heslop-Harrison, 1968a). It requires nexine to be formed (Lou et al., 2014), thus its formation follows the nexine formation. Its composition is most similar to type I primary cell walls, comprising cellulose, HG, RG-I, and AGPs (Geitmann et al., 1995; Ferguson et al., 1999; Drakakaki et al., 2006; Li et al., 2010; Heslop-Harrison, 1968a; Majewska-Sawka et al., 2004; Li et al., 1995; Cankar et al., 2014). In the absence of arabinan side-chains of RG-I, potato pollen grains collapse, highlighting their structural roles (Cankar et al., 2014). Similarly, a lack of arabinose for cell wall biosynthesis results in collapsed pollen grains due to defective intine (Drakakaki et al., 2006). Silencing of FASCICLIN-LIKE ARABINOGALACTAN PROTEIN3 (FLA3), a homologue of SOS5/FLA4, has defective intine and collapsed pollen 116  grains as a consequence (Li et al., 2010). These findings demonstrate that similar components are important for the structure of mucilage and intine.  Figure 4.3: Pollen wall structure of the mature pollen grain. Figure based on Ariizumi and Toriyama (2011) and Suzuki et al. (2008).  4.1.3 Objectives My main objective was to understand the breadth of biological functions of galactose oxidase-like proteins in plants. I took a reverse genetic approach, starting with a phylogenetic analysis and the analysis of the expression pattern of RUBY homologues, GALACTOSE OXIDASE-LIKE (GOXL) proteins, followed by a reverse-genetic analysis. The biochemical function of the enzymes was tested using transgene complementation of the ruby mutant. I present evidence that GOXL1 and GOXL6 are highly similar in sequence, have similar expression patterns and are biochemically functional equivalents of RUBY. In addition, I present evidence suggesting that GOXL1 and GOXL6 may have roles in pollen wall synthesis.   117  4.2 Results 4.2.1 Phylogenetic analysis of GOXL proteins from Arabidopsis The amino acid sequences of all seven genes from Arabidopsis thaliana that were annotated as “glyoxal oxidase-related” on TAIR (https://www.arabidopsis.org) were used to construct a phylogenetic tree. To eliminate the variability coming from the signal sequences, signal sequence predictions were made using SignalP 4.1 (http://www.cbs.dtu.dk/services/SignalP/) and Phobius (http://phobius.sbc.su.se/), and predicted signal sequences trimmed according to the most conservative prediction. A Maximum Likelihood tree was constructed using multiple alignment made with MAFFT programme.    118   Figure 4.4: Unrooted Maximum Likelihood phylogenetic tree of GOXL proteins from Arabidopsis thaliana compared with putative glyoxal oxidase from Vitis pseudoreticulata (VpGLOX) and characterised fungal galactose oxidases, alcohol oxidases and glyoxal oxidases (CAZy AA5 family). Node labels represent bootstrap support values based on 1000 replicates.  Plant enzymes belonging to the AA5 family cluster together and appear to be most closely related to fungal glyoxal oxidases (Figure 4.4), similar to what was observed for RUBY. The Arabidopsis members seem to have undergone a duplication, where GOXL3 is a putative paralogue of RUBY, GOXL1 of GOXL6, and GOXL4 of GOXL5. Based on grouping with a 119  putative GOXL from another plant species and highest distance from other AtGOXL members, GOXL2 is most likely to be the closest to the ancestral form. To better understand the evolution of GOXLs, a phylogenetic tree with GOXL orthologs from other angiosperms was constructed using Maximum Likelihood method. In the analysis, GOXL amino acid sequences from Capsella rubella were used to follow duplications in Brassicaceae family, sequences from Beta vulgaris to detect putative duplications within eudicots, and sequences from a basal angiosperm, Amborella trichopoda, to be able to follow the evolution of these proteins within angiosperms. From the tree, it is evident that GOXL3 and RUBY, as well as GOXL1 and GOXL6, resulted from a duplication within Brassicaceae family. This can be inferred from the observation that B. vulgaris sequences appear as sister clades to subclades in which each A. thaliana isoform groups with a single C. rubella isoform. This indicates that GOXL3/RUBY and GOXL1/GOXL6 are indeed paralogues. GOXL4 and GOXL5 do not seem to be recent paralogues, as GOXL4 and its orthologue from C. rubella form a clade with a B. vulgaris protein. This clade is a sister clade to a clade with GOXL5, suggesting that GOXL4 and GOXL5 separated before Brassicaceae. These findings suggest that RUBY and GOXL3, and GOXL1 and GOXL6, are more likely to be redundant than GOXL4 and GOXL5. 120   Figure 4.5: Unrooted Maximum Likelihood phylogenetic tree of angiosperm GOXL proteins. ‘ATR’ designates Amborella trichopoda sequences, ‘Bv’ Beta vulgaris sequences, and ‘Carubv’ Capsella rubella sequences. Node labels represent bootstrap support values based on 1000 replicates.  If GOXLs are functionally redundant, one would expect them to be present in the same subcellular compartment, i.e. the cell wall. Subcellular distribution of GOXLs was analysed using Aramemnon (http://aramemnon.uni-koeln.de/), a database which uses multiple transmembrane prediction programmes and generates consensus predictions. The consensus prediction score generated using Bayesian analysis of the results from multiple prediction programmes (AramLocCon score) indicates that all the members are most likely located in the secretory pathway (Table 4.1). However, cleavable signal peptide was not predicted for all proteins (Table 4.1), suggesting that only some are likely to be secreted to the apoplast through the endomembrane system, whereas others are either not secreted or secreted via alternative pathways. Another predictor of subcellular location of the proteins is the isoelectric point (pI). If 121  the protein is in an environment with a pH value close to its pI, it will lose charge and become insoluble (Arakawa and Timasheff, 1985). Proteins with high pI (RUBY, GOXL1, GOXL6 and GOXL5) would have a net positive charge in the apoplast, which is usually close to pH 5.5 (Cho et al., 2012; Li et al., 2005), and remain soluble. Since the difference between its pI and apoplastic pH is large, GOXL3 would probably behave the same. On the other hand, GOXL2 and GOXL4 are likely to be insoluble in the cell wall, which is inconsistent with the Aramemnon prediction for GOXL4. Therefore, direct experimental evidence is necessary to ascertain their subcellular location.  Table 4.1: Arabidopsis thaliana members of GALACTOSE OXIDASE-LIKE (GOXL) family of proteins with predicted subcellular location and signal peptide based on Aramemnon database. Isoelectric point predictions are taken from TAIR. Protein name Gene ID Predicted cleavable signal peptide Consensus prediction of secretory pathway location (AramLocCon) UniProt ID Predicted isoelectric point (pI) GOXL2 At3g53950 No 16.0 Q9M332 5.31 GOXL6 At5g19580 Yes 22.5 F4K172 9.93 GOXL1 At1g67290 Yes 26.2 Q9FYG4 9.29 GOXL3 At1g75620 No 15.7 Q9LR03 7.35 RUBY At1g19900 Yes 21.1 Q93Z02 9.03 GOXL4 At3g57620 Yes 25.7 Q9SVX6 6.19 GOXL5 At1g14430 No 24.3 Q9M9S1 8.92  4.2.2 Tissue-wide expression analysis of GOXL genes Publicly available expression data based on microarray experiments are not available for all the members of the AtGOXL family. Therefore, we used β-glucuronidase (GUS) reporter 122  system to investigate the expression of GOXLs. The entire non-coding regions upstream of GOXL sequences (Figure 4.6, grey) were used as promoters to drive GUS expression.   Figure 4.6: The structure of GOXL genes including the entire transcriptional regulatory region between the GOXL 5ʹ UTR and the nearest gene upstream. The position of the mutation found in various mutant alleles used in this study are marked with a black triangle (T-DNA or transposon insertion) or black arrow (insertions/deletions generated using CRISPR-Cas9 system).  Entire organs of T2 or T3 plants, coming from multiple independent T1 plants, were stained for GUS activity overnight to determine the expression domain driven by the promoter. 123  Once the activity was detected and found to be consistent between independently transformed lines, the incubation times were optimised to reduce overstaining and eliminate background. Consistent with the reports by Phan et al. (2011), GOXL1 expression was anther-specific and higher in the late developmental stages up to formation of mature pollen grains (Figure 4.7). When assays for GUS activity were longer (16 h), staining was very prominent in the anthers of older flowers (Figure 4.7). Individual flower buds at floral stages 11, 12 and 13 (Smyth et al., 1990) were dissected and incubated in the GUS reaction buffer for 3.5 hours to clearly distinguish whether the staining was derived from the gametophyte or the sporophyte. Such staining revealed that ProGOXL1 was most active at, or close to, anthesis (stage 13; Figure 4.7 K). A similar pattern was observed for ProGOXL6, with the exception that at stage 11, the staining was not observed in microspores, but in what appears to be the interlocular septum (Figure 4.7 F, black arrowhead). The activity of both promoters within the mature unfertilized flower seemed to be pollen-specific. The transmitting tract (septum) of 1 DPA elongating pistils/developing siliques did not stain, suggesting that the genes are not expressed in pollen tubes (Figure 4.7 N and O). Occasionally, stain appeared on the pistils following fertilization, but this was excess stain from overstained pollen grains that adhered to the surface of epidermal cells (Figure 4.7 O, arrows). These expression pattern similarities suggest that GOXL1 and GOXL6 may be redundant. 124   Figure 4.7: ProGOXL1 and ProGOXL6 are transcriptionally active late in pollen development. Wild type (Col-2; A, D, G, J, M) and representative lines for ProGOXL1:GUS (B, E, H, K, N) and ProGOXL6:GUS (C, F, I, L, O) were stained for GUS activity (blue colour). (A to C) Entire inflorescences stained overnight. Bars = 2 mm. (D to L) Dissected flowers stained for 3.5 hours. Bars = 200 μm. (M to O) 1 DPA pistils/siliques stained overnight. Bars = 500 μm. Black arrowhead indicates stained septum between locules. Black arrows indicate overstained pollen grains. 125   ProGOXL2 was found to be active in pistils of developing flowers (Figure 4.8 B). Prior to fertilization staining appeared to be uniform along the length of the pistil. Following anthesis staining became increasingly limited to the floral receptacle and stigma of the developing seed pod (Figure 4.8 B, D). The activity was also detected along the septum between the two silique valves (Figure 4.8 F, black arrow) and in the funiculus connecting the seed to the septum (Figure 4.8 D and F, black arrowhead).    126   Figure 4.8: ProGOXL2 is active in the developing pistil, silique septum and funiculi. Wild type (Col-2; A, C, E) and a representative line for ProGOXL2:GUS (B, D, F) stained for GUS activity overnight (blue colour). Black arrowheads indicate stained funiculus; black arrow indicates stained septum. Bars = 1 mm.  The staining pattern of ProGOXL3 lines appeared throughout the plant including the stem leaf and root, but it seemed to be limited to the vascular system. To pinpoint the exact location of the GUS activity, stems were sectioned at the immature (2 cm from the top), intermediate (5 cm 127  from the top) and mature (3 cm from the bottom) developmental stages of the stem vascular system. Staining was observed at all the stages in what appears to be phloem (Figure 4.9). The expression was also observed in leaves (Figure 4.9 F) and roots (Figure 4.9 H), demonstrating that it is not specific to the stem phloem.   128   Figure 4.9: ProGOXL3 is active in the phloem. Wild type (Col-2; A, C, E and G) and a representative line for ProGOXL3:GUS (B, D, F and H) stained for overnight for GUS activity (blue colour). (A) and (B) Inflorescence stem. Bars = 0.5 mm. (C) and (D) Transverse sections of the inflorescence stem. Bars = 100 μm. (D) Blue staining of the phloem. (E) and (F) Rosette leaves. Bars = 2 mm. (F) Weakly stained midrib of a leaf. (G) and (H) Seedlings with roots. Bars = 2 mm. (H) Blue staining of the stele in the centre of the root. 129  The staining of the entire inflorescences indicated that GUS driven by the ProGOXL4 was present in developing anthers but not mature anthers (Figure 4.10 B). When individual flowers were staged and stained for 3.5 hours, it was possible to identify GUS activity in the locules to what is most likely the tapetum (Figure 4.10 D). Similar to what was observed with the inflorescences, the activity was weaker in anthers of stage 12 flowers and it was absent in stage 13 flowers.   Figure 4.10: ProGOXL4 is active in the tapetum of developing anthers. Wild type (Col-2; A and C) and a representative line for ProGOXL4:GUS (B and D) stained for GUS activity (blue colour). (A) and (B) Inflorescences stained overnight. Bars = 1 mm. (C and D) Stage 11 flowers stained for 3.5 hours. Bars = 100 μm. 130   A putative promoter region of GOXL5 drove expression of GUS in all organs examined. GUS activity was detected in the midribs of leaves (Figure 4.11 D), and several layers of stem cortex (Figure 4.11 F). These observations suggest that GOXL5 is expressed in parenchyma.  Figure 4.11: ProGOXL5 is active in the parenchyma. Wild type (Col-2; A, C, and E) and a representative line for ProGOXL5:GUS (B, D, and F) stained for overnight for GUS activity (blue colour). (A) and (B) Inflorescences. Bars = 1 mm. (C) and (D) Rosette leaves. Bars = 1 mm. (D) Blue staining of the leaf midrib. (E) and (F) Transverse sections of the inflorescence stem. Bars = 200 μm. (F) Staining of the parenchyma in the stem cortex.   131  4.2.3 Functional complementation of ruby with GOXL genes In order to determine whether the GOXL homologues have a function similar to RUBY, each of the GOXL genes was tested for functional complementation of the ruby mutant phenotype. Coding sequences of GOXL genes were placed downstream of the ProRUBY, including the 5ʹ UTR, and sequences encoding the Citrine fluorescent protein were fused in frame to the 3ʹ end of the open reading frame. The constructs were transformed into mum2-1 ruby-1 plants and the seed mucilage phenotype of T2 seeds determined (Figure 4.12).  Figure 4.12: T2 seeds of ProRUBY:GOXL-Citrine in mum2-1 ruby-1 plants. Seeds were hydrated in water and stained with ruthenium red. Number of lines out of total lines displaying a phenotype shown in an image is written underneath. Bars = 200 μm.  132  All of the lines expressing GOXL1 under the control of ProRUBY, and all but one expressing GOXL6, displayed a mum2-like phenotype (Figure 4.12 E and L; Table 4.2), suggesting that these genes are able to complement ruby. Even though the extrusion was not observed, all lines had stained mucilage in the mucilage pockets, distinguishing them from mum2. Only a single transformant carrying ProRUBY:GOXL2-Citrine was recovered. This transformant displayed a ruby-like phenotype (Figure 4.12 F) suggesting a lack of complementation, but it is difficult to know if this one line is representative. Three out of five lines carrying ProRUBY:GOXL3-Citrine displayed partial suppression seen as an intermediate phenotype between a ruby-like and a mum2-like phenotype (Figure 4.12 H). Since RUBY is secreted and a signal sequence was not predicted to be present in the GOXL3 protein (Table 4.1), this was a surprising observation. GOXL4 driven by the ProRUBY was not able to complement ruby (Figure 4.12 I), which was expected based on the lack of a predicted signal peptide in GOXL4 (Table 4.1). In contrast, ProRUBY:GOXL5-Citrine was able to partially complement ruby in about 70% of the lines (Figure 4.12 J and K; Table 4.2), suggesting GOXL5 is a putative galactose oxidase. This result was expected given that GOXL5 has a predicted signal peptide and a pI similar to RUBY (Table 4.1).   133  Table 4.2: Results of functional complementation of ruby by GOXL genes based on ruthenium red staining after hydration in water. Construct mum2-like transformants ruby-like transformants Successfully complemented ProRUBY:GOXL1-Citrine 8 0 8/8 (100%) ProRUBY:GOXL2-Citrine 0 1 0/1 (0%) ProRUBY:GOXL3-Citrine 3 2 3/5 (60%) ProRUBY:GOXL4-Citrine 0 5 0/5 (0%) ProRUBY:GOXL5-Citrine 5 2 5/7 (71%) ProRUBY:GOXL6-Citrine 10 1 10/11 (91%)   The biochemical activity of the GOXL proteins was further examined using qualitative assays. Due to poor or no hydration of mum2 and mum2-like seeds, they were pre-incubated in diluted potassium hydrogencarbonate (KHCO3) to allow substrates to access the enzymes in the seed coat. KHCO3 was previously observed to release mucilage from mum2 and it has a lower pH than Na2CO3, thus it was predicted to be less likely to have adverse effects on enzymes. This apparent release of mucilage did enhance enzyme activity in mum2 seeds, consistent with this hypothesis. It was expected that like mum2 seeds, mum2 ruby seeds transformed with constructs that rescued the ruby phenotype would have GalOx activity. To test this, T2 seeds were incubated in a buffer containing TMB, D-galactose and HRP (Figure 4.13). Similar to experiments with wild-type seeds showing RUBY activity, the seeds of complemented lines were expected to display a blue colour when H2O2 generated by GalOx (GOXLs) is used by HRP to oxidise colourless TMB into a blue product. Consistent with the complementation results, the ProRUBY:GOXL3-Citrine lines with a mum2-like phenotype had GalOx activity (Figure 4.13 E, wells 2, 3 and 5), whereas those transformed lines exhibiting ruby-like phenotype did not (Figure 134  4.13 E, wells 1 and 4). This result suggests that GOXL3 is biochemically the most similar to RUBY, consistent with them being paralogues and suggests that, despite the apparent lack of a signal sequence, GOXL3 is secreted to the apoplast. Seeds expressing ProRUBY:GOXL1-Citrine and ProRUBY:GOXL6-Citrine displayed strong complementation with almost no mucilage extrusion, and were thus expected to exhibit strong GalOx activity. ProRUBY:GOXL1-Citrine seeds displayed at comparable rates to GOXL3 (Figure 4.13 C), but seeds expressing GOXL6-Citrine showed activity only after assaying for long periods of time (Figure 4.13 H). Consistent with the ruthenium red staining, ProRUBY:GOXL4-Citrine lines did not show GalOx activity. Surprisingly, ProRUBY:GOXL5-Citrine lines with mum2-like phenotype did not show GalOx activity.  It is possible that some enzymes (RUBY, GOXL1 and GOXL3) are more stable in the seed coat apoplast, whereas the others (GOXL5 and GOXL6) are less stable and more prone to degradation by apoplastic proteases that are present at the end of the seed coat development (Tsai et al., 2017). Another reason for low or no activity could be that the pH optimum of the enzymes is different from that of RUBY. The seeds of lines showing highest activity were tested for GalOx activity at pH 4.5, 5.0, 5.5, and 6.0. The activity was highest at pH 6 for all lines showing activity, and the lines showing no activity did not show it at lower pH, suggesting that the lack of activity is not due to the differences in pH optima. 135   Figure 4.13: HRP-TMB colourimetric assay demonstrating that galactose oxidase activity is restored in ProRUBY:GOXL-Citrine lines. Blue colour indicates oxidation of the colourless TMB substrate by HRP in the presence of H2O2. Assays were performed in the dark at room temperature, and images taken at 5 min (A), 33 min (C) and (E), and 6 h 30 min [(B), (D), (F), (G), and (H)] of reaction time.  T3 seeds of plants expressing GOXL-Citrine proteins were imaged in late development to inspect subcellular location of these proteins. Their subcellular distribution was compared to that of RUBY, which appears in the secondary cell wall of the columella (Figure 4.14 A, arrowhead) surrounding the cone-shaped cytoplasm (Figure 4.14 A, c).  GOXL1-Citrine and GOXL5-Citrine appeared intracellular (Figure 4.14 B and F, c), but also secreted to the columella (Figure 4.14 B and F, arrowhead). GOXL2-Citrine (Figure 4.14 C) was mostly intracellular (Figure 4.14 C, c) with some signal in columellae (Figure 4.14 C, arrowheads). Lines expressing GOXL3-Citrine 136  that displayed partial complementation showed similar signal distribution to RUBY-Citrine (compare Figure 4.14 A and D), whereas no signal was observed in lines that displayed no complementation. The same was true for GOXL6-Citrine (Figure 4.14 G). As predicted, GOXL4-Citrine did not appear in columella or middle lamella (Figure 4.14 E), suggesting that it was not secreted.   Figure 4.14: Subcellular distribution of GOXL-Citrine in the seed coat epidermis of seeds from senescing siliques. Pseudocoloured images are showing side view of epidermal cells. White arrowheads indicate signal in the columella; c = cytoplasm. Bars = 15 μm.  4.2.4 Reverse-genetic analysis of GOXL mutants Reverse genetics was used to investigate the biological role of the GOXL genes. Seeds of lines homozygous for mutations in each of the genes were ordered from a seed stock centre. The GOXL genes from each line were amplified and sequenced to confirm the position of the insertion (Figure 4.6, black triangles), and plants examined for phenotypes in those tissues known to express the genes. In goxl1, goxl4 and goxl6 lines the focus was on pollen defects, 137  which usually result in short siliques due to male sterility. Similarly, goxl2-1 was expected to affect seed set, which would have effect on silique size or opacity. Phloem defects were expected in goxl3-1, which would most likely have pleiotropic effect on the entire plant if the nutrient supply was compromised. Finally, goxl5 could have problems with mechanical support. However, none of the single mutants exhibited obvious phenotypes.  Since GOXL1 and GOXL6 are paralogs with a similar expression pattern, I reasoned that redundancy was very likely. Hence, I sought to generate a goxl1 goxl6 double mutant. The only available T-DNA insertional lines, goxl1-1 and goxl6-1, had normal transcript levels and insertions close to the end of the genes. The putative truncated mRNAs of these mutants were expected to encode all the predicted catalytic amino acids.  Thus, a CRISPR-Cas9 approach was employed to generate single nucleotide indels simultaneously in GOXL1 and GOXL6. Based on a shorter silique phenotype, a plant carrying a single bp (110ΔC) deletion in GOXL1 (goxl1-4; Figure 4.6, arrow) and a two bp insertion (233_234::AA) in GOXL6 (goxl6-5) was identified. Both of the mutations resulted in a premature STOP codon close to the 5ʹ end of the coding sequence. Homozygous double mutants could not be found in the T2 or T3 generations, a segregation pattern consistent with male-sterility of the double mutant. Further genetic analysis is required to confirm that these mutations are indeed responsible for the phenotypes observed.    138   Figure 4.15: The line carrying goxl1-4 and goxl6-5 mutations is partially male sterile due to collapsed pollen grains. (A) to (C) Wild type (Col-2); (D) to (F) putative mutant. (A) and (D) mature de-stained siliques demonstrating that mutant (D) has lower seed set than wild type (A). Bars = 0.5 cm. (B) and (E) dry pollen grains showing collapsed pollen grains (arrowheads) in the mutant (F). Bars = 40 μm. (C) and (F) Auramine O-stained pollen grains showing collapsed pollen grains (arrowheads) in the mutant (F). Bars = 20 μm.    Plants carrying both mutations (goxl1-4 and goxl6-5; heterozygous at at least one locus) developed shorter siliques than wild type. These siliques contained fewer seeds (Figure 4.15 D). An examination of anthers from these plants identified pollen grains that appeared collapsed and smaller in size, often attached to the grains normal in appearance (Figure 4.15 E, arrowheads). Pollen was next stained with Auramine O (Figure 4.15 C and F), a dye used to stain sporopollenin. The collapsed pollen grains (Figure 4.15 F, arrowheads) stained more intensely than the normal ones. The reticulate pattern of the exine was not changed in the collapsed grains, but it appeared that tecta were closer to each other than in normal grains, probably resulting in 139  more intense staining. This suggests that the phenotype is most likely due to changes in layers other than the exine. The expression pattern of GOXL1 and GOXL6 in late stages of flower development suggested that these genes may be involved in processes at the end of development. Similar to promoter-GUS experiments, developing anthers at floral stages 11 and 12 were examined. No difference was observed between wild type and mutant anthers at stage 11 or 12 (Figure 4.16 C), but at anthesis (stage 13) or later, collapsed pollen grains were observed in the double heterozygote (Figure 4.16 D. arrowheads). If the genetic analysis confirms that goxl1-4 and goxl6-5 are indeed responsible for these phenotypes, the presented result indicates that GOXL1 and GOXL6 may function at the end of development, possibly to strengthen the pollen wall before dehydration.  Figure 4.16: Phenotype of the putative mutant becomes apparent at the anthesis. De-stained and cleared anthers of wild type (A) and (C), and putative mutant (B) and (D) plants. Bars = 50 μm. 140  4.3  Discussion Even though well studied biochemically, prior to the discovery of RUBY, biological functions had not been identified for GalOx enzymes. In the previous chapter, I describe the first such enzyme studied in plants, RUBY, which functions specifically in the seed coat epidermis to cross-link RG-I and promote cell adhesion. To delve further into the exploration of biological functions of these enzymes, the phylogenetic relationships, expression patterns, and ability to complement the ruby-1 mutation were investigated for the entire family of glyoxal oxidase-related genes (GOXLs) from Arabidopsis. The expression patterns of these genes were diverse and, like RUBY, tissue specific. None of the single mutants displayed any obvious phenotypes in the tissues where the genes are expressed. Only a subset of the family was able to complement ruby. Below I describe what can be concluded about the function of each of the six homologues.  4.3.1 Some GOXL genes have undergone duplications and regulatory neofunctionalisation or subfunctionalisation Duplicated genes that are retained in the genome most commonly acquire a novel function (neofunctionalisation), or divide the ancestral function among each other (subfunctionalisation) (reviewed in Prince and Pickett, 2002). Four out of seven GOXLs are predicted to be secreted (RUBY, GOXL1, GOXL5 and GOXL6), whereas the other three (GOXL2, GOXL3 and GOXL4) should be intracellular. However, imaging of subcellular distribution of Citrine-tagged proteins suggests that all but GOXL4 seem to be at least in part apoplastic, as suggested by the signal in the columella or the middle lamella. To function on cell wall polysaccharides, the GalOx enzymes would have to be secreted or present in the secretory pathway where these polysaccharides are synthesised. Therefore, the intracellular GOXLs could be oxidising different 141  substrates and have different biological functions, suggesting a putative regulatory neofunctionalisation. For all the genes, the expression appears to be tissue-specific, with GOXL1 and GOXL6 expressed in pollen grains, RUBY in the seed coat epidermis, GOXL3 in the phloem, GOXL2 in the pistil, GOXL4 in the tapetum, and GOXL5 in the parenchyma. This highlights the regulatory neofunctionalisation or subfunctionalisation at the expression level. The exception would be GOXL1 and GOXL6, which, in addition to being recent paralogs, also exhibit very similar patterns of expression.   4.3.2 GOXLs likely have distinct biological functions When expression patterns, subcellular location predictions, and biochemical functions of Arabidopsis galactose oxidases are considered, predictions of putative biological functions can be made even in the absence of obvious mutant phenotypes.  4.3.2.1 GOXL2 GOXL2 is the most distantly related gene in the GOXL family. It is expressed in tissues of the developing pistils and siliques after fertilisation (Figure 4.8). Based on the expression in the pistil style, silique septum and funiculi, a putative function could be to guide pollen tubes through the transmitting tract by an unknown mechanism. However, since only one line expressing ProRUBY:GOXL2-Citrine was recovered, clear conclusions about the function of GOXL2 based on expression cannot be made. In addition, GOXL2 did not complement RUBY, but it does seem to be secreted to some extent (Figure 4.14 C). The GalOx activity was not observed in seeds, suggesting that either it has a distinct enzyme activity or its activity in the apoplast in relatively 142  low. Thus, it remains unclear whether it has a biochemical function similar to RUBY or not. The goxl2-1 lines displayed no visual phenotypes, thus there are no additional clues to the function of GOXL2. GOXL2 remains the most elusive member of the GOXL family.  4.3.2.2 GOXL3 GOXL3 appears to be a paralogue of RUBY (Figure 4.5) with a phloem-specific expression pattern (Figure 4.9). Despite only partial complementation of ruby in some of the ProRUBY:GOXL3-Citrine lines (Figure 4.12 H), the qualitative enzyme assay with seeds suggests GOXL3 is a putative galactose oxidase. The facts that GOXL3 has a similar subcellular distribution to RUBY and similar activity on the seed surface suggest the enzyme is extracellular. This is surprising, since there is no predicted signal sequence, a sequence that is normally required for secretion from the endomembrane system. It is possible that GOXL3 is secreted from the cytosol via a non-conventional secretory pathway. This type of secretion has been proposed when proteins without obvious signal peptides were found in the cell wall (Agrawal et al., 2010). Based on the promoter-GUS experiments, phloem is the tissue in which GOXL3 should be functioning. More intense staining at the top of the stem and at the tip of the root (Figure 4.9 H) suggests that the expression is strongest early in phloem development. The hypothetical function of GOXL3 based on the results is cross-linking RG-I in the cell walls of the phloem to promote cell adhesion and structural integrity of the tissue. Even though the information on phloem cell walls is scarce, RG-I-related galactans seem to be present in the Arabidopsis phloem (Torode et al., 2018), suggesting that a substrate for RUBY-like type of cross-linking is present. The goxl3-1 mutant had no obvious visual phenotype, but it is possible that a phenotype will only be obvious under specific physiological conditions. 143  4.3.2.3 GOXL4 GOXL4 was not able to complement the ruby mutation, which means that its biochemical function remains unknown. It does not appear to be secreted or function in the cell wall. This means that GOXL4 possibly underwent a regulatory neofunctionalisation. The expression of GOXL4 seems to be specific to the locules of the developing anthers, which is most likely the tapetum. Cells of the tapetum undergo programmed cell death in a process that involves reactive oxygen species (Xie et al., 2014). Based on the amino acid sequence similarity to AA5 family of enzymes, GOXL4 is likely an oxidase, thus able to generate H2O2 as a by-product of oxidation of an unknown substrate. Therefore, it may be one of the contributors to the ROS-mediated programmed cell death of the tapetum. Male sterility was not observed in the goxl4 mutants, thus it is hard to make a conclusion about its biological function.  4.3.2.4 GOXL5 GOXL5 seems to be, at least in part, functionally equivalent to RUBY, as suggested by partial complementation of the ruby phenotype by the ProRUBY:GOXL5-Citrine. This also suggests that GOXL5 is in the cell wall. GOXL5-expressing seeds did not show galactose oxidase activity, possibly due to instability that results in low protein levels in the columella and the middle lamella of mature seeds. The expression pattern suggests that GOXL5 has a function in parenchymatous cells (Figure 4.11). Since the expression was not detected in all of the parenchyma (e.g. stem pith), it seems that GOXL5 may only be expressed in the parenchyma that has roles in mechanical support. PECTIN METHYLESTERASE35 (PME35), functioning to de-esterify HG in the stem cortex, is important for mechanical support of the stem (Hongo et al., 2012), suggesting that pectin in the cortical parenchyma has roles in mechanical support. 144  Therefore, it is possible that GOXL5 functions to mediate cell-to-cell adhesion between cortical parenchyma, thus contributing to the mechanical support. The goxl5 mutants displayed no obvious phenotypes in stem tissues where it is expressed suggesting that if there is a phenotype, it may be conditional to a specific set of environmental conditions.  4.3.2.5 GOXL1 and GOXL6 possibly function to establish proper pollen wall structure Phylogenetic analysis indicated that GOXL1/GOXL6 have undergone a recent duplication and all existing data suggest that these two gene are completely redundant. The expression of these genes appears to be pollen-specific (Figure 4.6), and most prominent at later stages of development (stage 12 flowers) or at the time of the anthesis (stage 13 flowers). Previous reports on GOXL1 expression using promoter-GUS reporter system suggest that the expression is sporophytic and commences at anther developmental stage 8 (Phan et al., 2011), which corresponds to the floral development stage 10. This inconsistency may be due to the inclusion of a larger sequence upstream of GOXL1, including a part of the 3ʹ UTR of the adjacent gene, in the Phan et al. (2011) study, possibly containing additional cis-regulatory elements. Both GOXL1 and GOXL6 have GalOx activity on the seed surface, with GOXL6 showing lower activity possibly due to lower stability in the apoplast of dry seeds. Both genes have a predicted signal sequence, are expressed exclusively in anthers and complement the ruby mutation. Although neither single mutant appears to have a mutant phenotype, the CRISPR-Cas9-generated line carrying goxl1-4 and goxl6-5 exhibits partial sterility with shorter siliques and a lower seed set. This phenotype was correlated with 25% of the pollen grains appearing collapsed (Figure 4.15 E, F and G). In addition, no homozygote was recovered, suggesting a likely gametophytic defect consistent with GOXL1 and GOXL6 expression in pollen grains. Thus, a 145  gametophytic role in pollen development seems the most likely explanation for GOXL1/6 function. However, without further genetic analysis it is unclear whether goxl1-4 and goxl6-5 mutations are directly responsible for the phenotypes observed. Given its ability to complement a ruby mutation, putative extracellular location, and gametophytic function it is possible that GOXL1/6 are involved in the formation of the intine, the only part of the pollen wall fully produced by the gametophyte. Judging from the auramine O staining, the exine does not appear to be affected.  Support for this hypothesis comes from the literature, where defects in RG-I or HRGPs that result in defective intine and collapsed pollen grains have been reported (Cankar et al., 2014; Li et al., 2010). Since RG-I appears to have a role in structural support of the pollen grain through its side-chains, it is possible that GOXL1 and GOXL6 perform a function analogous to RUBY’s, where they oxidise Gal in the side-chains of RG-I to create hemiacetal cross-links. The timing of GOXL1 and GOXL6 expression at the end of pollen development when the tissue is undergoing maturation and dehydration (Heslop-Harrison, 1979) is consistent with such a role. Dehydration is regulated by the folding of the pollen wall at the apertures. In case of the tricolpate pollen produced by Arabidopsis, the pollen folds inwards at the apertures (colpi), covered by the water-permeable intine, and parts of the pollen grain covered by the interapertural exine close together to prevent further dehydration and collapse of the pollen grain in a process named harmomegathy (Figure 4.18; Katifori et al., 2010). In this way the pollen grain ends up fully covered by sporopollenin-rich exine, protecting the cell from environmental stresses and dehydration. It is, therefore, possible that GOXL1 and GOXL6 are necessary to properly establish the intine structure to allow proper folding or support for the interapertural exine during harmomegathy. Once this process is compromised by the absence of these proteins, 146  the apertures fail to fold properly, resulting in excessively desiccated and collapsed pollen grains (Figure 4.17).   Figure 4.17: Proposed harmomegathy-based model showing collapse of the pollen grain during desiccation in the putative goxl1 goxl6 mutant. In the wild type (left), the apertures (grey) fold in and allow buckling of the three colpi (yellow), sealing the pollen grain and preventing excessive desiccation. In the goxl1 goxl6 (right), incorrect folding results in collapsed pollen grains due to excessive desiccation.  147  Chapter 5: Conclusions 5.1 Galactose oxidase can cross-link plant cell walls In this dissertation I describe RUBY PARTICLES IN MUCILAGE (RUBY), the first GalOx with a known biological function, and the first GalOx studied in plants. RUBY is able to oxidize terminal Gal on sidechains of RGI and appears to cross-link pectin in cell walls of seed coat epidermal cells late in seed coat development. The primary function of this cross-linking is most likely to make cell-to-cell adhesion stronger between adjacent seed coat epidermal cells, as well as between seed coat epidermal cells and the underlying palisade cells to resist the strong shear forces generated by mucilage extrusion. GalOx enzymes have been shown to cross-link polysaccharides in vitro, resulting in gels that become insoluble after dehydration (Mikkonen et al., 2014). The insolubility is the result of hemiacetal cross-links that form between C-6 aldehydes of oxidised Gal and hydroxy groups of proximal polysaccharides. My data are consistent with the hypothesis that RUBY promotes similar covalent bonds within the pectin of cell walls.    5.1.1 Future directions My thesis research has generated a number of questions that remain unanswered relating to plant galactose oxidases and their roles in vivo.  First, direct evidence of hemiacetal existence in the cell walls of seed coat epidermal cells is still lacking. Hemiacetals in cell walls could be detected using Fourier-transform infrared (FT-IR) spectroscopy or nuclear magnetic resonance (NMR) spectroscopy. For instance, if mum2 ruby is compared to mum2, it would be possible to look for differences in the spectra between the two and see if the difference may be interpreted as a hemiacetal. However, the difficulty with this approach would be observing the native state in 148  muro. The cross-linking prevents extrusion and mucilage would have to be extracted using a base to break hemiacetals, followed by dehydration to re-form them.  A second unresolved question is the exact substrate for RUBY in the middle lamella. The branched RG-I released in ruby mutant mucilage might represent that substrate. However, the branched RG-I lacks terminal Gal and instead has terminal Ara branches on each Gal sidechain. It is unclear whether the terminal Ara is present in wild type or if it is unique to ruby mucilage.  It is possible that wild-type branched RG-I exists with at least some terminal Gal sidechains and therefore is a substrate for RUBY.  A way to test if wild-type seeds produce RG-I without Ara is to look at the monosaccharide composition of the seeds during development. If Ara branches are added onto RG-I in response to ruby mutation, the difference is expected to be seen only after RUBY expression. In this situation, wild type is expected to have lower Ara than ruby once RUBY is expressed (9-10 DPA). Another possibility is that Ara branches are removed in wild type by unknown RUBY-dependent arabinosidases. In this case, similar levels of Ara would be present in both wild-type and ruby seeds prior to RUBY expression. After RUBY is expressed, Ara may be cleaved only in wild type (lower Ara), but not in ruby (higher Ara). If no difference between the two is observed, then it is likely that branched RG-I is not the substrate.   5.2 Putative galactose oxidases in Arabidopsis may function in specialised tissues In Chapter 4, I characterised the family of putative galactose oxidases of Arabidopsis thaliana using phylogenetic, molecular genetic, biochemical, and cytological studies. The members of this family have been annotated as glyoxal oxidase-related, in accordance with the amino acid sequence comparisons and phylogenetic analyses. All members of Arabidopsis glyoxal oxidase-related (GOXL) family lack tryptophan (Trp) found in the active site of typical GalOx enzymes, 149  but not glyoxal oxidases (Whittaker et al., 1999). Despite the absence of Trp in the active site, RUBY, GOXL1, GOXL3, GOXL6, and possibly GOXL5, are able to influence galactose oxidase activity in seeds. Wild-type seeds, expressing RUBY, displayed a higher rate of glycerol oxidation than D-galactose oxidation, which is consistent with the observations that the absence of indole side-chain of Trp in FgGalOx, and replacement with a non-branched glycine (G), allows glycerol to access the active site (Rogers et al., 2007). This indicates that Trp is not necessary for GalOx activity, but due to the amino acid sequence differences, it cannot be excluded that another Trp or similar amino acid from the sequence takes its place in the active site of Arabidopsis enzymes. The biological roles of GOXL enzymes are unknown, but some clues come from expression results and a putative goxl1 goxl6 mutant plant. Highly tissue-specific expression patterns of each gene suggest that they probably function in specialised tissues. RUBY’s likely paralogue, GOXL3, seems to have undergone regulatory neofunctionalisation or subfunctionalisation, suggested by the expression pattern in the seed coat in case of RUBY, and phloem in case of GOXL3. As discussed in Chapter 4, GOXL3 may be functioning on galactans, rich in the cell walls of phloem sieve elements, to reinforce the pectin of cell walls beyond that the Ca2+-cross-linking of HG. The additional support of the cell wall may be necessary in sieve elements, as they are experiencing constant changes in osmotic pressure caused by loading and unloading of sucrose (Gould et al., 2005). GOXL2 seems to be the closest to the ancestral form of GOXLs. The functional characterisation of this protein was unsuccessful, but the expression analysis suggests that it should be functioning in the pistil. GOXL4 is most closely related to GOXL5, but they are not recent paralogues and are expected to have diverged more than RUBY/GOXL3 in terms of expression. GOXL4 seems to be intracellular and expressed in the 150  tapetum of developing anthers, whereas GOXL5 is extracellular and expressed in parenchymatous cells throughout the plant. As there is no evidence for enzymatic function of GOXL4, it is hard to speculate about its potential roles in the tapetum. GOXL5 is able to partially complement ruby, but it showed no galactose oxidase activity in an enzyme assay, making it only a putative GalOx. If it is functionally similar to RUBY, it may be promoting RG-I cross-linking between the cells of the parenchyma with roles in mechanical support (e.g. stem cortex). As the goxl5 mutants have no obvious phenotypes, this type of cross-linking may be an additional support to the Ca2+-cross-linking between HG in the cortex, a process which seems to be necessary to keep the stem upright (Hongo et al., 2012). GOXL1 and GOXL6 are similar in sequence and appear to be paralogues with similar expression patterns, suggesting likely genetic redundancy. They are both expressed in pollen grains late in the development, but their transcription does not remain active in the pollen tubes. Double mutants were generated to overcome likely redundancy, and plants with mutations in both GOXL1 (goxl1-4) and GOXL6 (goxl6-5) were isolated based on putative partial male sterility. These plants have collapsed pollen grains and shorter siliques with lower number of seeds, but further genetic analysis is needed to confirm that the goxl1-4 and goxl6-5 are responsible for these phenotypes. If they are, it would suggest that GOXL1/6 are important in establishing the pollen wall structure late in development. Since the phenotype was observed at anthesis when pollen dehydration occurs, it is possible that gametophytically-expressed GOXL1/6 cross-link RG-I in the intine. Similar to the function of RUBY in the drying seed coat, oxidised RG-I can cross-link to intine polysaccharides through hemiacetals to prevent misfolding of the apertures during controlled dehydration (harmomegathy).  151  5.2.1 Future directions The biological functions of GOXL enzymes other than RUBY are still unknown due to the lack of genetic evidence. The pollen phenotypes suggest that GOXL1/6 may have roles in strengthening pollen wall, but genetic analysis needs to be completed to confirm that mutations are segregating with the phenotypes. This can be addressed through observations of segregation after a cross to wild type, or through complementation of mutant plants using a wild-type copy of the genes. If the modification of the intine in the putative mutant occurs at the end of development, no difference is expected to be observed in the intines of wild type and goxl1 goxl6 mutants prior to anthesis. Therefore, transmitting electron microscopy may be used to inspect the intine of the developing pollen to test this hypothesis. Determining if harmomegathy is indeed the process affected can be done by growing flowers in high humidity, which may prevent dehydration leading to collapse, resulting in normal, yet hydrated, pollen grains in the putative mutant.  Further investigation of GOXL2 function would require more extensive experimentation, starting from exploring whether it actually has galactose oxidase function by testing the activity in vitro. Even though the phenotype is not obvious in goxl2 line, further exploration of the timing and exact location of the expression of GOXL2 may give clues as to when and how to look for phenotypes.  GOXL3 is a putative galactose oxidase with an unknown biological function. Distinguishing the exact cell type of the phloem in which GOXL3 is expressed would be the first step towards forming hypotheses about its biological function. It is possible that the goxl3 only exhibits phenotypes under certain conditions, such as changes in osmotic pressure in the sieve 152  element, thus examining the mutant under different environmental conditions, e.g. varying day/night cycles, may be the next step in exploring the functions of this gene.  Due to probable intracellular distribution of GOXL4, and inability to complement ruby, it may be necessary to demonstrate that this protein has galactose oxidase activity. Following this would be understanding the nature of the isolated goxl4-1 and goxl4-4 mutants. Even though homozygotes were isolated, they exhibited no obvious phenotype that a mutant defective in tapetal function would exhibit, such as male sterility. It is possible that truncated proteins, if they are expressed at all, are still able to perform their function. Another possibility is that the phenotypes of these mutants are too subtle to be observed.  Even though direct biochemical evidence for the galactose oxidase function of GOXL5 is lacking, partial complementation of ruby suggests that this enzyme may have a similar enzymatic function to RUBY. If the enzyme is more prone to protease degradation in the seed coat apoplast, it may be necessary to attempt the expression in heterologous systems, such as Nicotiana benthamiana or Pichia pastoris, to demonstrate its activity in vitro. Homozygous goxl5 mutants exhibited no stem phenotypes. As suggested before, HG cross-linking may be sufficient in cross-linking the middle lamellae of cortex parenchyma during normal conditions, thus obscuring the role of hemiacetal cross-links. Making a cross between goxl5 and pme35 may be a good way to test this hypothesis. If GOXL5 has an effect on this process, the goxl5 pme35 is expected to have more severe phenotype than pme35 or goxl5 alone. Goxl5 phenotypes may have to be investigated using compression tests of the stem, similar to what was to done to explore phenotypes of pme35 (Hongo et al., 2012). Atomic force microscopy can also be used to test the cell wall and middle lamella strength in stem sections to detect possible subtle phenotypes. 153  5.3 Summary The research presented in this thesis sheds new light on galactose oxidases by identifying both biochemical and biological functions, and suggests a novel type of cell wall cross-linking involving hemiacetals. 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